U.S. patent number 8,748,091 [Application Number 12/971,240] was granted by the patent office on 2014-06-10 for characterizing stretched polynucleotides in a synthetic nanopassage.
This patent grant is currently assigned to The Board of Trustees of the University of Illinois, The John Hopkins University. The grantee listed for this patent is Aleksei Aksimentiev, Jeffrey Comer, Utkur Mirsaidov, Gregory Timp, Winston Timp. Invention is credited to Aleksei Aksimentiev, Jeffrey Comer, Utkur Mirsaidov, Gregory Timp, Winston Timp.
United States Patent |
8,748,091 |
Timp , et al. |
June 10, 2014 |
**Please see images for:
( Certificate of Correction ) ** |
Characterizing stretched polynucleotides in a synthetic
nanopassage
Abstract
Methods of trapping a deformed portion of a double-stranded
polynucleotide in a membrane nanopassage are provided. In an
aspect, the membrane has a nanopassage that defines a confine
region, wherein the membrane separates a first fluid compartment
from a second fluid compartment, and the nanopassage is in fluid
communication with the first and second compartments. A
polynucleotide is provided to the first fluid compartment and
optionally a threshold voltage for the membrane and the
polynucleotide is determined. A driving voltage across the membrane
that is greater than the threshold voltage is applied to force a
portion of the polynucleotide sequence into the nanopassage confine
region, and decreased to a holding voltage bias to trap the
polynucleotide portion in the nanopassage confine region. In
particular, at least one nucleotide base-pair is fixably positioned
in the nanopassage confine volume. In further embodiments, any of
the trapping methods are used to characterize or sequence double
stranded DNA.
Inventors: |
Timp; Gregory (South Bend,
IN), Timp; Winston (Baltimore, MD), Mirsaidov; Utkur
(Urbana, IL), Aksimentiev; Aleksei (Urbana, IL), Comer;
Jeffrey (Urbana, IL) |
Applicant: |
Name |
City |
State |
Country |
Type |
Timp; Gregory
Timp; Winston
Mirsaidov; Utkur
Aksimentiev; Aleksei
Comer; Jeffrey |
South Bend
Baltimore
Urbana
Urbana
Urbana |
IN
MD
IL
IL
IL |
US
US
US
US
US |
|
|
Assignee: |
The Board of Trustees of the
University of Illinois (Urbana, IL)
The John Hopkins University (Baltimore, MD)
|
Family
ID: |
44646353 |
Appl.
No.: |
12/971,240 |
Filed: |
December 17, 2010 |
Prior Publication Data
|
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|
|
Document
Identifier |
Publication Date |
|
US 20110226623 A1 |
Sep 22, 2011 |
|
Related U.S. Patent Documents
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Application
Number |
Filing Date |
Patent Number |
Issue Date |
|
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61287974 |
Dec 18, 2009 |
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Current U.S.
Class: |
435/6.1; 205/543;
435/283.1; 536/23.1; 422/82.05 |
Current CPC
Class: |
G01N
33/48721 (20130101); G01N 27/4146 (20130101); C12Q
1/6869 (20130101); B01L 3/5027 (20130101) |
Current International
Class: |
C12Q
1/68 (20060101); C12M 1/36 (20060101); G01N
27/00 (20060101); C07H 21/04 (20060101) |
References Cited
[Referenced By]
U.S. Patent Documents
Foreign Patent Documents
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WO2008/079169 |
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Jul 2008 |
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WO |
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WO2009/030953 |
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Mar 2009 |
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WO |
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WO 2010/080617 |
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Jul 2010 |
|
WO |
|
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|
Primary Examiner: Forman; Betty
Attorney, Agent or Firm: Lathrop & Gage LLP
Government Interests
STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT
This invention was made with United States government support under
contract number R01 HG003713 awarded by the National Institute of
Health and contract number PHY0822613 awarded by the National
Science Foundation. The United States government has certain rights
in the invention.
Claims
We claim:
1. A method of characterizing at least a portion of a
double-stranded polynucleotide, said method comprising: providing a
membrane having a nanopassage that defines a confine region,
wherein said membrane separates a first fluid compartment from a
second fluid compartment, and said nanopassage is in fluid
communication with the first and second compartments; providing
said double-stranded polynucleotide to said first fluid
compartment; establishing a driving voltage bias that is greater
than a threshold voltage across said membrane to force a portion of
said double-stranded polynucleotide into said nanopassage, wherein
said portion of said double-stranded polynucleotide has a confined
portion positioned within said confine region and said confine
region deforms said double-stranded polynucleotide by increasing
axial rise from an undeformed axial rise value to a deformed axial
rise value in said confine region or a region adjacent thereto;
monitoring an electrical current through said nanopassage;
identifying a confine state from said a monitored electrical
current from said monitoring step, wherein said confine state
corresponds to said confined portion containing a confined
nucleotide base-pair of said polynucleotide in said confine region;
reducing said driving voltage bias to a holding voltage that is
less than or equal to said threshold voltage, thereby trapping said
confined nucleotide base-pair in said nanopassage confine region
for a trapping time, wherein said trapping time is selected from a
range that is greater than or equal to 10 ns and less than or equal
to 1 second; and measuring a nucleotide base-pair dependent current
blockade through said nanopassage confine region having said
confined nucleotide base-pair during said trapping time, to
characterize the confined nucleotide base-pair, thereby
characterizing at least a portion of said polynucleotide.
2. The method of claim 1, further comprising: establishing a
translocation voltage bias that is greater than the threshold
voltage to force said confined nucleotide base-pair out of said
confine region in a direction that is toward said second
compartment; repeating said monitoring, identifying, reducing and
measuring steps to thereby characterize a confined nucleotide
base-pair that is at a position upstream from the previously
characterized confined nucleotide.
3. The method of claim 2, wherein the characterized confined
nucleotides base-pair are nucleotide base-pairs adjacent to each
other in said polynucleotide.
4. The method of claim 2, further comprising repeating the steps of
claim 2 to characterize a contiguous portion of said
polynucleotide, wherein said contiguous portion corresponds to at
least 10% of the entire length of said polynucleotide.
5. The method of claim 4, wherein said contiguous portion
corresponds to the entire length of said polynucleotide.
6. The method of claim 2, wherein said translocation voltage is
less than said driving voltage bias.
7. The method of claim 2, wherein said holding voltage is applied
for a holding time sufficient to provide high-fidelity assessment
of said confined nucleotide base-pair.
8. The method of claim 2 wherein said polynucleotide travels in a
direction from said first compartment to said second compartment at
a translocation velocity that is greater than or equal to 1
nucleotide per 10 nanoseconds or greater than or equal to 1
nucleotide base pair per 10 nanoseconds.
9. The method of claim 2, further comprising: diagnosing a medical
condition for a patient from which the polynucleotide is
obtained.
10. The method of claim 9 wherein the medical condition relates to
a specific polynucleotide sequence.
11. The method of claim 9, wherein the medical condition relates to
a methylation parameter.
12. The method of claim 2, wherein the translocation voltage bias
is a voltage pulse having a duration that is less than or equal to
1 .mu.s.
13. The method of claim 2, wherein the magnitude of said
translocation voltage bias is at least two times greater than said
holding voltage.
14. The method of claim 1, wherein said polynucleotide translocates
unidirectionally from said first compartment to said second
compartment.
15. The method of claim 1, further comprising determining said
threshold voltage for said nanopassage and said polynucleotide.
16. The method of claim 1, wherein said characterizing is one or
more of identifying a nucleotide-type, a nucleotide base-pair type
or nucleotide methylation state.
17. The method of claim 1 wherein said characterization is
methylation content, methylation pattern, or methylation content
and pattern.
18. The method of claim 1, wherein said characterization is
determining at least a portion of said polynucleotide sequence.
19. The method of claim 1 wherein said deformed axial rise value is
at least 20% greater than the undeformed axial rise value.
20. The method of claim 1, wherein said deformed axial rise value
is selected from a range that is greater than or equal to 0.34 nm
and less than or equal to 0.7 nm.
21. The method of claim 1, wherein said characterization comprises
determining the sequence of at least 2000 contiguous bases.
22. The method of claim 1, wherein said confine region has a
minimum cross-sectional area that is less than or equal to 4.4
nm.sup.2.
23. The method of claim 22, wherein said nanopassage is a tapered
nanopore having a maximum diameter that is less than or equal to
2.6 nm and a minimum diameter centered in said confine region that
is selected from a range that is greater than or equal to 1 nm and
less than or equal to 2.4 nm.
24. The method of claim 1, wherein said membrane is a
Si.sub.3N.sub.4 membrane having a thickness selected from a range
that is greater than or equal to 5 nm and less than or equal to 30
nm.
25. The method of claim 1, wherein one nucleotide of said confined
nucleotide base-pair is uniquely identified with one strand of said
double stranded polynucleotide.
26. The method of claim 1, further comprising the step of obtaining
a high fidelity measure of the nucleotide base-pair dependent
current blockade by measuring the nucleotide base-pair dependent
current blockade over the trapping time and calculating an average
nucleotide base-pair dependent current blockade over the trapping
time.
27. A method of trapping a portion of a double-stranded
polynucleotide in a membrane nanopassage, said method comprising
the steps of: providing a membrane having a nanopassage that
defines a confine region, wherein said membrane separates a first
fluid compartment from a second fluid compartment, and said
nanopassage is in fluid communication with the first and second
compartments; providing said double-stranded polynucleotide to said
first fluid compartment; determining a threshold voltage for said
membrane and said double-stranded polynucleotide; establishing a
driving voltage bias across said membrane that is greater than said
threshold voltage, to force a portion of said double-stranded
polynucleotide into said nanopassage confine region, wherein said
portion of said double-stranded nucleotide in said confine region
is deformed and said confine region deforms said double-stranded
polynucleotide by increasing axial rise from an undeformed axial
rise value to a deformed axial rise value in said confine region or
a region adjacent thereto; and decreasing said driving voltage bias
to a holding voltage bias, wherein said holding voltage bias is
less than said threshold voltage, thereby trapping said
polynucleotide portion in said nanopassage confine region for a
trapping time, wherein said trapping time is selected from a range
that is greater than or equal to 10 ns and less than or equal to 1
second, wherein at least one nucleotide base-pair is fixably
positioned in said nanopassage confine volume during said trapping
time.
28. The method of claim 27, further comprising: measuring a
blockade current through said nanopassage having at least one
nucleotide base-pair positioned in said confine volume;
sequentially forcing said polynucleotide through said confine
volume nucleotide base-pair by nucleotide base-pair by switching an
electric field from a translocation voltage bias that is greater
than said threshold voltage to said holding voltage bias at a
switching frequency, wherein said holding voltage bias is applied
when a nucleotide base-pair is positioned in said confine region,
and said sequentially forcing step provides a nucleotide base-pair
stepwise movement of said polynucleotide through said confine
region in a direction from said first compartment to said second
compartment so that every nucleotide base-pair within a contiguous
length of said polynucleotide is trapped in said confine region and
said blockade current is measured for each trapped nucleotide
base-pair.
29. The method of claim 28, wherein the holding voltage bias
corresponds to no voltage difference across said membrane.
30. The method of claim 27, wherein said holding voltage is applied
for a holding time that is sufficient to provide high-fidelity
measurement of said blockade current for said nucleotide base-pair
positioned in said confine volume.
31. The method of claim 27, wherein said nanopassage confine region
that traps said portion of polynucleotide has a minimum
cross-sectional area that is 2.56 nm.sup.2.
32. The method of claim 27, wherein said polynucleotide is DNA
having a length that is greater than or equal to 200 base
pairs.
33. The method of claim 27, wherein six or fewer base pairs are
trapped in said nanopassage interior volume.
34. The method of claim 27, wherein the nanopassage is a pore
having a minimum diameter that is smaller than an average diameter
of said polynucleotide that is trapped.
35. The method of claim 27, further comprising measuring an
electrical blockade current across said nanopassage for said
trapped portion.
36. A method of sequencing a double-stranded polynucleotide, said
method comprising: providing a membrane having a nanopassage that
defines a confine region having a minimum dimension that is less
than an average axial diameter of said double-stranded
polynucleotide, wherein said membrane separates a first fluid
compartment from a second fluid compartment, and said nanopassage
is in fluid communication with the first and second compartments;
providing said double-stranded polynucleotide to said first fluid
compartment; establishing a driving voltage bias that is greater
than a threshold voltage across said membrane to force a portion of
said double-stranded polynucleotide into said nanopassage, wherein
said portion of said double-stranded polynucleotide has a confined
portion positioned within said confine region; monitoring an
electrical current through said nanopassage; identifying said
confined portion as a confined nucleotide base-pair from a
monitored electrical current from said monitoring step; reducing
said driving voltage bias to a holding voltage that is less than or
equal to said threshold voltage, thereby trapping said confined
nucleotide base-pair in said nanopassage confine region for a
trapping time, wherein said trapping time is selected from a range
that is greater than or equal to 10 ns and less than or equal to 1
second; measuring a nucleotide-dependent current blockade through
said nanopassage confine region having said confined nucleotide
base-pair during said trapping time, to identify the confined
nucleotide base-pair, thereby characterizing at least a portion of
said polynucleotide; establishing a translocation voltage bias that
is greater than the threshold voltage to translocate said
polynucleotide in a direction that is toward said second
compartment, wherein said translocation moves said polynucleotide
by one base-pair through the confine region; repeating said
monitoring, identifying, reducing and measuring steps to thereby
identify a confined nucleotide base-pair that is at a single
base-pair sequential position difference from the previously
characterized confined nucleotide base-pair, thereby sequencing at
least a portion of said double-stranded polynucleotide.
37. The method of claim 36 wherein the measuring step is repeated
over the entire polynucleotide length, thereby sequencing the
entire polynucleotide.
Description
CROSS-REFERENCE TO RELATED APPLICATIONS
This application claims benefit of U.S. provisional patent
application 61/287,974 filed Dec. 18, 2009 which is hereby
incorporated by reference to the extent it is not inconsistent with
the present disclosure.
BACKGROUND OF THE INVENTION
Provided are methods and devices for characterizing
polynucleotides, such as determining nucleotide sequences in double
stranded DNA.
DNA, the program for life, is encoded using four chemical bases
called adenine (A), guanine (G), cytosine (C) and thymine (T),
which are paired together in a complementary fashion (A to T and C
to G) and ordered in a species-specific sequence. The aim of
genomic science is to predict biological behavior using the
information stored in the DNA sequence within each cell. But when
the first draft sequence of the human genome emerged in early
2001,6 despite its enormous value to genetics, it quickly became
apparent that our understanding of the relationship between the
genetic code and cellular function was deficient. For example, only
5% of the human genome is conserved, and of that, only 30% lies
within the exons of known protein-encoding genes.7 The rest lies in
the so-called "dark matter" of the human genome--leading to efforts
such as the ENCyclopedia Of DNA Elements (ENCODE)8 that strive to
identify regulatory components. Identifying genes and controlling
regions, such as promoter sites turns out to be a major undertaking
in itself.
To glean more information about how genetics informs cellular
function, and therefore its affect on development and disease, it
is essential that we learn to sequence rapidly and economically
using a minute amount of material. The tasks can be categorized as
follows. On one hand, in terms of de novo sequencing, not only are
there many species with unsequenced genomes, but the human
microbiome, the genome sequence of the many different species of
bacteria living in or on humans, still remains a mystery. The
microbiome of the flora of our gut alone is estimated to contain
.about.300 billion base-pairs (Gbp), or .about.100.times. the human
genome.9 On the other hand, the vast majority of the work ahead
involves re-sequencing genomes with an already known base sequence.
The first, obvious example is mutation sequencing where recent work
has shown that the majority of human cancers do not always have
mutations in the same locations, or even the same genes.10
Moreover, the mutations and genotype of the individual has been
shown to be important for chemotherapeutic effectiveness: i.e.
genomics can determine a drug's effectiveness on an
individual.11
Sequencing can also provide clues to health and development beyond
the actual genomic sequence itself. The proteins expressed by genes
represent the machinery of the cell--they make things work. But an
individual organism can express the same genes differently
depending on the epigenetic profile. High-throughput sequencing
aspires to determine this profile. For example, it can give
information on DNA-binding protein interaction, using ChIP-seq to
find the locations of occupied binding sites.12 With inexpensive,
high-throughput sequencing, we will be able to determine the
difference between these binding sites in different tissue and
under different conditions. Moreover, using ChIP-seq we can also
achieve single-base resolution of the genomic histone code, one of
the epigenetic regulators of chromatin structure and gene
expression.13 It can also be used to determine DNA methylation
patterns, a reversible modification of cytosines (in mammals),
which alters protein binding (see, e.g., U.S. Pat App. No.
61/139,056 hereby incorporated by ref.) Subsequent sequencing and
alignment may be used to distinguish methylated from unmethylated
cytosines, illuminating the methylation pattern.14 It may also be
advantageous for gene expression studies to sequence the
transcriptome; i.e. the sequence of the RNA extracted from cells or
tissues. This can give detailed information about the levels of
expression, the splicing variation, and even allow for the
identification of new non-coding RNAs, which may be involved in the
regulation and are parts of the "dark matter" of the genome.15
All of these "-omes" would be facilitated by technology that
inexpensively and quickly determines sequence information from a
genetic sample. For this reason, ultra-low cost sequencing
technologies have been identified as a scientific priority and
significant effort is being devoted to its study and
development.
Since its development in 1977, the Sanger method of DNA sequencing
has transformed biology--it is the standard to which all other
methods of sequencing are compared. 16 The basis for Sanger
sequencing is the polymerase chain reaction (PCR), which is used in
combination with dideoxy-terminated nucleotides triphosphates to
prematurely terminate the elongation reaction. The classical
chain-termination method requires a DNA template, a DNA primer, a
DNA polymerase, nucleotides and fluorescently labeled nucleotides
that terminate DNA strand elongation. By mixing fluorescently
labeled dideoxynucleotides with deoxynucleotides, the PCR reaction
is prematurely terminated, leading to fragmentary single stranded
copies of the template that differ in length with the last base
fluorescently labeled with a different fluorescent moiety,
depending on the base. Separating these fragments by size through
electrophoresis, the sequence can be determined from the color of
fluorescence produced at a given length.
Though functional, this procedure is problematic for several
reasons. The template read length using this method is limited to
.about.800 bp. This introduces significant challenges, especially
for de novo sequencing, requiring that either chromosome walking or
shotgun sequencing be used, which are both time consuming and
require re-assembly of the completed sequence. The chain
termination reaction is also time consuming, as is electrophoretic
separation, leading to the development of techniques for massively
parallel methods for sequencing.17 But the overarching problems
with Sanger sequencing method are the relatively large amounts of
DNA required--amplification leads to errors--and the expense due to
reagents for labeling and separation.
There are emerging technologies that have the potential to
supersede conventional, Sanger sequencing and in some cases
sequence the human genome for $1000 or less. Shendure et al have
analyzed these technologies in detail.18 They can be loosely
categorized as: bioMEMs, which is just an extension of conventional
electrophoretic methods through miniaturization and integration;
sequencing-by-hybridization, which uses the differential
hybridization of oligonucleotide probes to decode the DNA sequence;
massively parallel signature sequencing (MPSS), which is based on
cycles of restriction digestion and ligation; and finally,
non-enzymatic, real-time single-molecule sequencing.
BioMEMs has the advantage that it relies on the same tested
principles as electrophoretic sequencing, which has already been
used to sequence 10 11 nucleotides. Using variations of the Sanger
process in conjunction with capillary array electrophoresis to
separate deoxyribonucleotide triphosphate fragments, about 100 bp
can be sequenced per minute at a cost of <$1 with an accuracy of
about 99.99%, which is considered to be the gold standard, but it
seems unlikely that a factor of 100,000.times. cost reduction will
be achieved through scaling and integration alone. Hybridization
sequencing has the advantage that the data collection method, i.e.
scanning the florescence emitted by labeled DNA that has been
hybridized to an array of probe sequences, is compatible with
high-throughput, but probes have to be designed that avoid
cross-hybridization to the wrong target. This renders 50% of the
chromosome inaccessible. All methods like sequence-by-synthesis,
cyclic-array sequencing on amplified molecules, and MPSS, which
rely on some method of isolated clonal amplification are, first of
all, costly and often problematic. For example, they may experience
a low frequency of nucleotide misincorporation or
non-incorporation, which manifests itself in signal decay through
"dephasing". In contrast, cyclic-array sequencing on single
molecules eliminates the costly PCR-amplification step, requires
less starting material with no risk of de-phasing, but achieving
the signal-to-noise required for single molecule detection is still
a challenge.
According to Mardis,19 right now the Roche GS-FLX (454) sequencer
uses emulsion PCR to produce 100 Mb of data in 7 h with a 250 bp
read length (per bead) at a cost of $8439 or $84.40 per Mb. In
contrast, a run in an Applied Biosystems SOLiD (sequencing by oligo
ligation) sequencer requires 5 days and produces 3-4 Gb of sequence
data with an average read length of 25-35 bp, costing $5.81 per Mb.
Applied Biosystems estimates that their SOLiD sequencer will be
able to sequence an entire human genome for only $10,000 in just 2
weeks. Following Shendure's analysis, 18 for re-sequencing, the
error rate has to be less than the expected variation in the
sequence. Since human chromosomes differ at approximately 1 in
every 1000 bp, an error rate of 1/100 kbp would be needed to ensure
confidence. If the accuracy of a raw read is 99.7% (current
state-of-the-art), then .times.3 coverage of each base will yield
this error rate. To ensure a minimum .times.3 cover of >95% of
the diploid human genome, .times.6.5 coverage, or about 40 billion
raw bases at a cost per base of <$10000, or 4 million bases per
$1. If an improvement over SOLiD performance is derived simply from
an increase in the acquisition rate per device, we would therefore
need to sequence at a rate of .about.330,000 bp/s to reach a $1,000
genome. No assembly is required in re-sequencing a genome; the read
needs only be long enough to allow it to be matched to a unique
location in an assembled reference genome, and how it differs from
the reference. In the mammalian genome only .about.73% of 20-bp
genomic reads SOLID uses can be assigned to a single unique
location. Achieving >95% uniqueness will require reads >60
bp. Thus, a re-sequencing instrument that can deliver a $1,000
human genome with reasonable coverage and accuracy will need to
achieve >60 bp reads with 99.7% raw-base accuracy, acquiring
data at a rate of 330,000 bp/s or 1 bp/3 .mu.s. A faster instrument
with longer reads will be cheaper still.
Single molecule DNA sequencing represents the logical
end-of-the-line in development of sequencing technology, which
extracts the maximum amount of information from a minimum of
material and pre-processing. When paired with a high-throughput and
low cost instrument, it would change the genomic flow of data from
a trickle to a deluge. Specifically, the low material requirement
coupled with quick results would allow for easy sequencing of
precious primary samples from human patients, e.g. allowing doctors
to sequence a biopsy from a tumor to determine the best
chemotherapy. Moreover, it would represent a leap forward in
determining the epigenome, the non-genetic marks on DNA which
affect gene expression.
SUMMARY OF THE INVENTION
Provided are methods, devices, and methods for making devices
capable of sequencing DNA via confined electric field control
across a nanopassage in a membrane, such as a nanopore, through
which a polynucleotide is forced to traverse. Sequencing a single
molecule of DNA with a nanopore is a revolutionary change in
sequencing technology because it combines the potential for long
read lengths (>5 kbp) with high speed (1 bp/10 ns), while
obviating the need for costly procedures like PCR amplification or
sample preparation due to the exquisite single molecule
sensitivity. Moreover, electrical detection of DNA using a
nanopassage could have several advantages over cyclic arrays or
fluorescent microscopy. Usually single molecule sequencing relies
on enzymatic incorporation of a fluorescently labeled
mononucleotide through a polymerase and applying techniques that
suppress the ambient radiation so that one molecule can be
identified. In contrast, the nanopassage sequencing concept uses a
radically new approach to detection that is reminiscent of
Coulter's original idea of using objects within a constricted
current path to alter, the electrical resistance.
Nanopore sequencing relies on the electric signal that develops
when DNA translocates through a pore in a membrane. DNA is a highly
charged polyanion. By applying an electric field to a
nanometer-diameter pore in a thin membrane, individual DNA
molecules are forced to move through the pore in a single-file
sequential order, as if threading a needle. Because of unique
composition of each base-type, each base has a characteristic
electrical signature. Assessing the electrical signature over time
as a single polynucleotide traverses a pore having a confined
electric field provides a means for analyzing nucleotide sequence.
In this manner, a pore can by used to analyze the sequence by
reporting all of the signatures in a single read without resorting
to multiple copies of DNA.
The methods provided herein provide the capability to reliably
sequence a polynucleotide by a unique trapping method to ensure
there is a sufficient residence time, on a base-by-base basis, as a
polynucleotide traverses a nanopassage. The combination of a
time-controlled electric field across a nanopassage having a
well-defined geometry, results in a functional equivalent of a
harmonic trap. The nanopassage is designed to provide at least a
portion of the passage, such as a confine region, with a dimension
sufficiently small such that the polynucleotide cannot traverse and
transit without undergoing a deformation. In particular, when a
nucleotide base or nucleotide base pair is forced into the confine
region by an applied electric field that deforms the
polynucleotide, lowering the electric field means the portion of
polynucleotide within the confine region cannot exit the confine
region, and is trapped. This trap means that electric measurements
may be reliably made, effectively for as long as desired, to
provide high-fidelity measurement on a specific region, including a
specific base, of a DNA molecule. The electric field is switched to
a sufficiently high magnitude to permit the trapped portion to
deform and exit the confine region, and when the next base or base
pair is in the confine region, the electric field is lowered to
again trap the polynucleotide, but at a different axial
position.
In an aspect, provided are methods of trapping a polynucleotide or
sequencing a polynucleotide in a nanomembrane nanopassage. In an
aspect, any of the methods provided herein related to a
polynucleotide that is double stranded, such as double stranded
DNA.
In an embodiment, provided is a method of characterizing at least a
portion of a double-stranded polynucleotide by providing a membrane
having a nanopassage that defines a confine region. The membrane
separates a first fluid compartment from a second fluid
compartment, and the nanopassage is in fluid communication with the
first and second compartments. A polynucleotide is provided to the
first fluid compartment. A driving voltage bias that is greater
than a threshold voltage is established across the membrane to
force a portion of the polynucleotide sequence into the
nanopassage, wherein the polynucleotide portion has a confined
portion positioned within the confine region and the confine region
deforms the double-stranded polynucleotide structure by increasing
axial rise from an undeformed axial rise value to a deformed axial
rise value in the confine region or a region adjacent thereto.
Electrical current is monitored through the nanopassage and a
confine state identified from the monitored electrical current,
wherein the confine state corresponds to the confined portion
containing a confined nucleotide base-pair of the polynucleotide in
the confine region. The driving voltage bias is reduced to a
holding voltage that is less than or equal to the threshold voltage
and that is greater than or equal to zero, thereby trapping the
confined nucleotide base-pair in the nanopassage confine region for
a trapping time. During the trapping, a nucleotide base-pair
dependent current blockade is measured through the nanopassage
confine region having the confined nucleotide base-pair, to
characterize the confined nucleotide base-pair, thereby
characterizing at least a portion of the polynucleotide.
In an embodiment, the method further comprises establishing a
translocation voltage bias that is greater than the threshold
voltage to force the confined nucleotide base-pair out of the
confine region in a direction that is toward the second
compartment. The monitoring, identifying, reducing and measuring
steps are repeated to thereby characterize a confined nucleotide
base-pair that is at a position upstream from the previously
characterized confined nucleotide. For example, the characterized
confined nucleotides base-pair can be nucleotide base-pairs
adjacent to each other in said polynucleotide. In this manner, the
characterization may be continuous polynucleotide base-pair or base
sequencing. In an aspect, the method relates to characterizing a
contiguous portion of the polynucleotide, wherein the contiguous
portion corresponds to at least 10% of the entire length of the
polynucleotide. In another aspect, the contiguous portion
corresponds to the entire length of the polynucleotide, or at least
90%, 95% or 99% of the entire length. In an aspect, the entire
length of the polynucleotide is sequenced.
In an embodiment, the method relates to a polynucleotide that
translocates unidirectionally from the first compartment to the
second compartment.
In another aspect, the method further comprises determining the
threshold voltage for the nanopassage and the polynucleotide. The
threshold determination may be empirical, such as by monitoring
electric current through the nanopassage under different electric
field conditions to identify when the polynucleotide is capable of
translocating and when the polynucleotide is reliably trapped over
a desired trapping time, as described herein.
In an aspect, the characterization is one or more of identifying a
nucleotide-type, a nucleotide base-pair type or nucleotide
methylation state. In an aspect the nucleotide is a
naturally-occurring nucleotide. The method, however, is compatible
for any type of nucleotide, so long as the nucleotide provides an
electrically-detectable signal that is distinct from other
nucleotides sought to be differentiated.
In an aspect, the characterization is methylation content,
methylation pattern, or methylation content and pattern, such as
further described in 61/139,056. In another aspect, the
characterization is determining at least a portion of the
polynucleotide sequence.
The method is optionally further described in terms of a deformed
axial rise value, such as a value that is at least 20% greater than
the undeformed axial rise value. In an aspect, the undeformed axial
rise is about 0.34 nm per base-pair or less, corresponding to the
double helix of double stranded DNA so that the deformed axial rise
is about 0.4 nm or greater. In an aspect, the deformed axial rise
value is selected from a range that is greater than or equal to
0.34 nm and less than or equal to 0.7 nm. In general, the axial
rise is sufficiently small such that there is not physical
permanent damage to the polynucleotide.
In another embodiment, the characterization comprises determining
the sequence of at least 2000 contiguous bases. The methods and
systems provided herein are compatible with sequencing a wide range
of polynucleotide lengths, including lengths that are less than
2000 bases, such as by manipulating membrane capacitance. For
example, capacitance values ranging from about 10 to 400 pF are
used in various embodiments. In general, the larger the capacitance
the slower the electrical response in the system, as characterized
from the associated time constant for the system (e.g., the product
of the electrolyte resistance and membrane capacitance) compared to
the translocation time (e.g., product of the polynucleotide length
and translocation velocity). Contemplated herein is miniaturization
to significantly reduce membrane capacitance, thereby decreasing
the minimum detectable strand length. For example, the time
constant may be reduced substantially (such as to values less than
1 ns), such as by the use of composite membranes made of
Si.sub.3N.sub.4 and polyimide.
In an aspect, the invention is further described in terms of the
nanopassage. For example, a nanopassage having a confine region
with a maximum cross-sectional area that is less than or equal to
the cross-sectional area of hydrated double stranded DNA. In an
aspect the maximum cross-sectional is defined by two axes, which
are both comparable to or less than 2.9 nm in length, since the
hydrated diameter of double-stranded DNA ranges from 2.6 to 2.9 nm.
Thus, the maximum cross-sectional area is 8.41 nm.sup.2. A
nanopassage may also have a confined region with a minimum
cross-sectional area that is greater than or equal 1.6.times.1.6
nm.sup.2. For an area less than this the double-stranded DNA
molecule will denature and translocate through the pore one strand
at a time. (40).
In an aspect the nanopassage is a pore having a diameter that is
less than or equal to the effective diameter of a hydrated DNA
double helix. In an aspect, the nanopassage is a tapered nanopore
having a maximum diameter that is less than or equal to 2.9 nm and
a minimum diameter centered in said confine region that is selected
from a range that is greater than or equal to 1.6 nm and less than
or equal to 2.9 nm. In an aspect, the taper relates to a maximum
diameter at a membrane surface and a minimum diameter in the
central portion of the membrane. In an aspect, the minimum
dimension is at the midpoint between the top and bottom surfaces of
the membrane and the taper is linear and can be described by a
taper angle. In an aspect, the taper angle is selected from a range
that is greater than or equal to 10.degree. and less than or equal
to 30.degree.. In an aspect, the membrane has a thickness that is
greater than or equal to 5 nm and less than or equal to 100 nm.
In an embodiment, the process relates to trapping the
polynucleotide at a desired base-pair for a trapping time. In an
aspect, the trapping time is selected from a range that is greater
than or equal to 10 ns and less than or equal to 60 second. In an
aspect, the trapping time is for a sufficiently long time to
provide a high-fidelity read of the current blockade through the
nanopassage for the confined base-pair. Accordingly, in an aspect,
the holding voltage is applied for a holding time sufficient to
provide high-fidelity assessment of said confined nucleotide
base-pair. "High fidelity" refers to a measurement related to a
specific base or base-pair that is sufficiently reliable that it is
capable of being statistically distinguished from a measurement
related to a different specific base or base-pair. For example, for
measurements that are noisy, prolonging the measurement time may
provide a more reliable average value and reduce the associated
standard deviation, thereby providing a measurement of sufficient
fidelity, e.g., high fidelity, to provide a statistically
significant difference with a measurement obtained for a different
base or base-pair. "High fidelity" also refers to the ability to
correctly identify the base or base-pair more than 95% of the time,
such as by providing a p-value of 0.05 at the 95% confidence
level.
In an aspect, the translocation voltage is less than the driving
voltage bias.
In an aspect, the polynucleotide travels in a direction from the
first compartment to the second compartment at a translocation
velocity that is greater than or equal to 1 nucleotide per 10
nanoseconds or greater than or equal to 1 nucleotide base pair per
10 nanoseconds.
In an embodiment, any of the methods provided herein further relate
to diagnosing a medical condition for a patient from whom the
polynucleotide is obtained, such as a medical condition that
relates to a specific polynucleotide sequence or to methylation
parameter/profile.
In an embodiment, the invention relates to one or more
characteristics of the translocation voltage bias. In an aspect,
the translocation voltage bias is a voltage pulse having a duration
that is less than or equal to 1 .mu.s. In an aspect the magnitude
of the translocation voltage bias is at least two times greater
than said holding voltage.
In an aspect, the membrane is a Si.sub.3N.sub.4 membrane, such as
SiN membrane having a thickness selected from a range that is
greater than or equal to 5 nm and less than or equal to 30 nm.
In another embodiment the invention is a method of trapping a
portion of a double-stranded polynucleotide in a membrane
nanopassage by providing a membrane having a nanopassage that
defines a confine region, wherein the membrane separates a first
fluid compartment from a second fluid compartment, and the
nanopassage is in fluid communication with the first and second
compartments. A polynucleotide is provided to the first fluid
compartment and a threshold voltage for the membrane and the
polynucleotide is determined. A driving voltage bias is established
across the membrane that is greater than the threshold voltage, to
force a portion of said polynucleotide sequence into the
nanopassage confine region, wherein the nucleotide portion in the
confine region is deformed and the confine region deforms the
double-stranded polynucleotide structure by increasing axial rise
from an undeformed axial rise value to a deformed axial rise value
in said confine region or a region adjacent thereto. The driving
voltage bias is decreased to a holding voltage bias, wherein the
holding voltage bias is less than the threshold voltage, thereby
trapping the polynucleotide portion in the nanopassage confine
region for a trapping time, wherein at least one nucleotide
base-pair is fixably positioned in the nanopassage confine
volume.
In an aspect the threshold voltage is determined empirically. In an
aspect the threshold voltage is determined by referring to a
specification supplied by the manufacturer, such as depending on
specific membrane composition and configuration, nanopassage size
or geometry, DNA characteristic (e.g., length), electrolyte
composition, electrode geometry, etc.
In an aspect, the method further comprises measuring a blockade
current through the nanopassage having at least one nucleotide
base-pair positioned in the confine volume and sequentially forcing
the polynucleotide through the confine volume nucleotide base-pair
by nucleotide base-pair by switching an electric field from a
translocation voltage bias that is greater than the threshold
voltage to the holding voltage bias at a switching frequency. In
this manner, the holding voltage bias is applied when a nucleotide
base-pair is positioned in the confine region, and the sequentially
forcing step provides a nucleotide base-pair stepwise movement of
the polynucleotide through the confine region in a direction from
the first compartment to the second compartment so that every
nucleotide base-pair within a contiguous length of the
polynucleotide is trapped in the confine region and the blockade
current is measured for each trapped nucleotide base-pair.
In an aspect, the holding voltage is applied for a holding time
that is sufficient to provide high-fidelity measurement of the
blockade current for the nucleotide base-pair positioned in the
confine volume. In an aspect, the holding voltage bias corresponds
to no voltage difference across the membrane. Alternatively, the
holding voltage is an AC voltage. Alternatively, the holding
voltage is a small positive or a small negative bias.
In an embodiment, the nanopassage confine region that traps the
portion of polynucleotide has a maximum cross-sectional area that
is 8.41 nm.sup.2 and a minimum cross-sectional area of 2.56
nm.sup.2. In an aspect, the polynucleotide is DNA, such as double
stranded DNA forming a double helix, having a length, such as a
length that is greater than or equal to 200 base pairs or shorter
lengths.
In an embodiment, six or fewer base pairs are trapped in the
nanopassage interior volume, or in the confine region.
In an aspect, the nanopassage is a pore having a minimum diameter
that is smaller than an average diameter of said polynucleotide
that is trapped. In an aspect, the nanopassage is a tapered pore
having a maximum diameter at one surface and a minimum diameter
positioned between the two membrane surfaces or at the opposite
surface of the membrane. In an aspect, the taper is from each
membrane surface and meet in between the surfaces, or at about the
middle between the two surfaces.
In an aspect, the method relates to measuring an electrical
blockade current across the nanopassage for said trapped
portion.
In another embodiment, the invention is a method of sequencing a
double-stranded polynucleotide. In an aspect, a membrane is
provided having a nanopassage that defines a confine region having
a minimum dimension that is less than an average axial diameter of
the hydrated double-stranded polynucleotide, wherein the membrane
separates a first fluid compartment from a second fluid
compartment, and the nanopassage is in fluid communication with the
first and second compartments. A polynucleotide is provided to the
first fluid compartment. A driving voltage bias that is greater
than a threshold voltage is established across the membrane to
force a portion of the polynucleotide sequence into the
nanopassage, wherein the polynucleotide portion has a confined
portion positioned within the confine region. An electrical current
through the nanopassage is monitored and the confined portion is
identified as a confined nucleotide base-pair from the monitored
electrical current. The driving voltage bias is reduced to a
holding voltage that is less than or equal to the threshold voltage
and that is greater than or equal to zero, thereby trapping the
confined nucleotide base-pair in the nanopassage confine region for
a trapping time. A nucleotide-dependent current blockade is
measured through the nanopassage confine region having the confined
nucleotide base-pair, to identify the confined nucleotide
base-pair, thereby characterizing at least a portion of the
polynucleotide. A translocation voltage bias is established that is
greater than the threshold voltage to translocate the
polynucleotide in a direction that is toward the second
compartment, wherein the translocation moves the polynucleotide by
one base-pair through the confine region. The monitoring,
identifying, reducing and measuring steps are sequentially repeated
to thereby identify a confined nucleotide base-pair that is at a
single base-pair sequential position difference from the previously
characterized confined nucleotide base-pair.
In an aspect, any of the methods provided herein relates to
repeating the measuring step over the entire polynucleotide length,
thereby sequencing the entire polynucleotide. In an aspect, any of
the methods provided herein relates to uniquely identifying one
nucleotide of the confined nucleotide base-pair with one strand of
said double stranded polynucleotide, such as assigning a base to
either the 5' or 3' strand.
Without wishing to be bound by any particular theory, there can be
discussion herein of beliefs or understandings of underlying
principles or mechanisms relating to embodiments of the invention.
It is recognized that regardless of the ultimate correctness of any
explanation or hypothesis, an embodiment of the invention can
nonetheless be operative and useful.
BRIEF DESCRIPTION OF THE DRAWINGS
FIG. 1: Sequencing dsDNA with a solid-state nanopore showing a
sequence of polynucleotide interactions (a)-(g) with a nanopore and
corresponding applied voltage (h) and detected current (i) through
the nanopore. (a) The driving bias of V.sub.0, such as
V.sub.0>V.sub.threshold(V.sub.T) causes a flow of ionic current,
I.sub.0 through the pore. (b) Eventually, the driving bias forces a
molecule of DNA into the pore opening, the molecule stretches and
threads the pore (c). Consequently, the ionic current through the
pore is at least partially blocked. The molecule is trapped by
lowering the driving voltage to V.sub.1<V.sub.T. (d) Using a
series of voltage pulses, the DNA advances through the pore. The
current is modulated distinctly by each base-pair (see FIG. 8). (e)
The blockade current is measured using low noise, phase-sensitive
lock-in techniques. (f-g) shows repeating of the cycle. (h) Diagram
of the bias to the membrane by the Ag/AgCl electrodes immersed in
the solution during each step in the process. (i) Diagram of the
ionic pore current. Bottom panel is a schematic of one embodiment
of a nanopassage and corresponding confine region, with respect to
double stranded DNA.
FIG. 2: (a) Membranes are formed by depositing a Si3N4 layer onto a
silicon substrate. An SEM cross-section through the membrane
structure is shown on the right. (b) DUV lithography and a
combination of dry and wet etching through the backside of the
wafer reveal the membrane. Subsequently, a photosensitive polyimide
layer on the front surface is deposited and patterned to reduce the
stray capacitance. An optical micrograph of a 10 .mu.m window in
polyimide used to define the membrane area is shown on the right.
(c) After revealing the membrane, a pore is sputtered in it using a
tightly focused, high energy electron beam. A nanopore through the
membrane is shown on the right. (d) An array of 1.8 nm nanopores
sputtered back-to-back in different membranes using similar
conditions.
FIG. 3: a) A TEM micrograph of a 2.5.times.2.0 nm nanopore in a
silicon nitride membrane 15 nm thick. (b) Electrolytic current
measured in 100 mM KCl at 800 mV (top trace) and 200 mV (bottom
trace) through the pore shown in (a) as a function of time. The
frequency of blockades decreases dramatically with voltage; at 200
mV no transients are observed, suggesting no translocation through
the nanopore and that the threshold voltage is greater than 200 mV.
(c) Three examples of current blockades observed in the pore shown
in (a) as a function of time at V=800 mV under the same conditions
as (b). The open pore current at this voltage is about 2.85 nA.
These current blockades are associated with .lamda.-DNA
translocating through the pore. (d) The frequency of blockades
observed at 800 mV with a particular change in current normalized
to the open pore current in the same pore. (e) The frequency of
blockades observed with the same pore as a function of membrane
voltage illustrating the frequency drop as voltage decreases below
0.5V. The dotted line represents a fit to the data. (f)
Distributions illustrating the frequency as a function of the
duration of a current blockade, t.sub.D, above threshold at 1.0 V
(smallest x-intercept), 800 mV (middle x-intercept) and 700 mV
(largest x-intercept). The distribution depends sensitively on the
voltage. Inset: tD-1, as a function of the applied voltage. (g)
Magnitude of the pore current as a function of frequency measured
through 3 different membranes: one with a nitride layer 12 nm
thick; another with a 200 nm thick nitride layer and the third with
a 30 nm thick nitride layer with a polyimide coating along with
corresponding fits to the data (solid lines) (h) Schematics of the
lumped element model for a composite polyimide/nitride membrane
with a nanopore in it. The model is superimposed on the physical
geometry (not to scale) and used to analyze the frequency responses
shown in (a). The Faradic impedances, double-layer capacitances,
depletion capacitance and series resistances are all represented
and fit using ADS. (i) Simplified lumped element model derived from
(h) (from Smeets (31)).
FIG. 4: (a) Noise power spectra of 3 nanopores measured in FIG.
3(g) with different effective capacitance. From bottom to top, a
300M.OMEGA. resistor, a 5 nm pore in polyimide covered
Si.sub.3N.sub.4 membrane, a 4 nm pore in .about.200 nm
Si.sub.3N.sub.4 membrane, and a 3 nm pore in 12 nm Si.sub.3N.sub.4
membrane. The low frequency 1/f noise (red line), the high
frequency dielectric noise (cyan line) along with the amplifier
noise (green) are analyzed for a 5 nm pore in polyimide coated
membrane. The fit to the total noise is shown in orange. (b) The
rms current noise as a function of bandwidth. The capacitive noise
predominates at high frequency. (c) The rms-current noise increases
with effective capacitance. (d) Effect of electrolyte concentration
on noise power spectra for a 3 nm pore in 12 nm thick nitride. 1/f
noise depends on electrolyte concentration while dielectric noise
does not. (e) .lamda.-DNA current blockade through two 3.0.+-.0.2
nm diameter pores: one in a 30 nm thick nitride membrane (top
trace) and another in the same membrane that is also coated with
3.6 .mu.m thick polyimide layer (bottom trace) except for a 10
.mu.m window. The peak-to-peak noise is dramatically improved with
polyimide.
FIG. 5: (a) Triggered by the onset of a blockade in the 2.5 nm
pore, the voltage across the pore is switched from 800 mV (above
the threshold for stretching) to 200 mV (below threshold). As a
result, the duration of the current transient (blue) increases from
about 200 .mu.s to about 6.73 s. The transient observed at 7.709 s
reflects a single DNA nucleotide base or base-pair exiting the
pore. The grey trace represents data taken at a 10 kHz bandwidth;
the blue trace is the same data with a 1 kHz (8-pole Bessel)
filter. Inset: Detail showing the onset of a current transient and
the 200 .mu.s built-in delay executed before triggering the voltage
step. (b) A magnified view of the pore current observed during
(left) and after (right) the blockade, showing the magnitude of the
current fluctuations. The blockade fluctuations are related to bps
translocating at 1 bp/221 .mu.s. (c) Distribution of the current as
a function of .DELTA.I/I0 during the blockade (left) and the open
pore (right). (d) The distribution of dwell times observed at 800
mV (grey), and after the voltage is switch from 800 mV to 200 mV to
the return to I0 (blue).
FIG. 6: Trapping a .lamda.-DNA molecule in a nanopore. (a) A 56 s
current blockade. Triggered by the onset of a blockade in a 2.5 nm
pore, the membrane voltage is switched from 600 mV (above
threshold, V.sub.T) to 100 mV (below threshold, V.sub.T). As a
result, the width of the current transient (blue) increases from
about 900 .mu.s to .about.56 s. The transient observed at 58.8 s is
supposed to indicate a .lamda.-DNA molecule exiting the pore. The
grey trace represents data taken at 10 kHz bandwidth; the blue
trace is the same data with a 1 kHz filter. (b) Distribution of the
current during the blockade for t<58.8 s observed in the
interval 40.0-40.5 s (left), and the open pore for t>58.8 s
(right) The distribution for the trapped molecule can be fit to two
Gaussians: one (solid blue) offset from the median (.DELTA.I=0) by
+2.86 pA with a width of 7.6 pA; and another (solid red line)
offset by -6.51 pA with a width of 5.8 pA. The black line
represents the sum. (c) Signal from base-pairs translocating
through an open pore. (d) Histograms showing the distribution of
dwell times observed at 600 mV (left-most histogram) and the
distribution of elapsed time spanning the instant when a blockade
event triggers the voltage switch from 600 mV to 150 mV (blue) or
100 mV (red) to the return of the current to the open pore value
seconds later. The peak in the distribution of current blockade
durations (blue) increases from about 900 .mu.s to about 200 ms,
increasing .times.200.
FIG. 7: MD simulation of the nanopore trap. (a) Snapshot of the
simulated system that includes dsDNA, a 2.6 nm.times.2.1 nm-cross
section pore, water and ions (not shown) at a transmembrane bias of
250 mV. The molecular conformation is stretched in the constriction
beyond the 0.34 nm about 8-20%. (b) The number of basepairs
permeating through this pore in four MD simulations carried out
applying different biases. The simulations predict a voltage
threshold for restarting the translocation between 500 mV and 1.0V.
(Inset) Histogram of the displacements of the basepair nearest to
the center of the membrane in a simulation carried out at a 0V
bias. The solid line shows the distribution for a harmonic trap
with a k=3.0.+-.0.8 nN/nm spring constant.
FIG. 8: Sequencing dsDNA by measuring the ionic current. (a) A
snapshot of dsDNA trapped in a 2.0-nm-diameter pore. The DNA
preserved its canonical B-form structure outside the pore
constriction, whereas in the constriction it is stretched. (b)
Snapshot of a simulation system used to determine the sensitivity
of the ionic current blockade to the type and orientation of the
confined basepair. A phantom nanopore of a 1.9 nm diameter and a
0.1M KCl solution is used for these simulations. (c) The simulated
values of the ionic current for each basepair system under an
effective bias of 750 mV with a pore current, I0=195.0.+-.5.2 pA
(estimated from a 717.4 ns simulation). The error bars show the
standard errors of the average currents. The total simulation time
for each system is specified in the bottom row. (d) Table of
absolute valies of the current computed at 0.1M and 1.4M KCl salt
concentrations.
FIG. 9 Streptavidin bound biotin DNA duplex trapped by the electric
field in a nanopore. (a) Model of biotinylated dsDNA bound to
streptavidin in a 2.6 nm.times.2.1 nm cross-section pore in a 23 nm
thick membrane. The molecular conformation is stretched in the
constriction beyond the 0.34 nm about 8-20%, depending on the
applied voltage. (b) The dependence of the peak dwell time for both
C-G and A-T duplexes on the membrane voltage at high salt, 1M KCl.
(c) The distribution of dwell times inferred from the blockade
current for both C-G and A-T duplexes measured for streptavidin
bound biotin DNA at 1V. The residence time in the pore is
extraordinarily long even at high voltage. (d,e,f) Current
blockades measured at 1V, 0.8V and 0.2V transmembrane bias in 1M
KCl in a 2.6.times.2.1 nm pore associated with C-G and A-T duplexes
biotinylated to streptavidin. The difference in blockade current
can be used to discriminate C-G from A-T. (g). Summary of the
difference between the open pore current and the blockade current
measured for C-G and A-T duplexes at 1M KCl.
FIG. 10 shows the dependence of the coefficient A measured in a
pore with a 2.4.+-.0.2 nm diameter in a 12 nm thick nitride
membrane.
FIG. 11 (a) The number of base-pairs permeating through a nanopore
(2.0 nm diameter) (compare against the larger 2.6 nm.times.2.1
nm-cross section pore of FIG. 7(b)) in four MD simulations carried
out at different biases. The simulations predict a voltage
threshold between 0.5V and 1.0V. (Inset) Histogram of the
displacement of the base-pair nearest to the membrane's center at a
0V bias. The solid line shows the distribution expected for a
harmonic trap with a 6.6 nN/nm spring constant. (b) Stepwise
advancement of dsDNA by a single base-pair in a 2 nm pore. The
position of the nucleotides near the constriction are plotted
versus time. In these simulations, a half-sine voltage pulse was
applied from 0.0 to 0.2 ns with amplitudes of 6 and 8 V. At 6 V
there is a small displacement of the nucleotide during the
application of the pulse, after which the nucleotide returns to its
initial position. However, at 8V the molecule translates so that
the final position of the nucleotides near the center of the pore
are near the initial positions of the nucleotide directly below,
constituting a nucleotide step.
FIG. 12 Sub-millisecond simulations of ionic current blockades
using atomically precise Brownian Dynamics (BD). The simulated
ionic current using microsecond all-atom and 100-microsecond BD
trajectories is platted as a function of base-pair type, with X-Y
and Y-X indicating bases on different strands. mC-G denotes
methylated cytosine. The much greater efficiency of the BD
simulations allows for much longer runs and therefore much smaller
error bars at a fraction of the computational cost.
FIG. 13: Magnified view of blockade current measured near 58.8 s
superimposed on a current trace comprised of the simulated current
response estimated from the results in FIG. 12 for the last 50 bp
of the 3' end of .lamda.-DNA. The correlation coefficient between
the measurement and the simulation is 0.31.
FIG. 14(a): Current blockades associated with .lamda.-DNA measured
at 1V in a high salt concentration (1M KCl) in a 2.6.times.2.1 nm
cross-section pore in a 23 nm thick membrane. Near 0.6 s the DNA
sticks to the pore. (b,c) Single molecule fluorescence of
.lamda.-DNA intercalated with YOYO fluorescent dye in high salt (1M
KCl). The DNA is absorbed on silica (b), but not on silicon nitride
membranes (c).
DETAILED DESCRIPTION OF THE INVENTION
As used herein, "double stranded" refers to two complementary
polynucleotide strands having base pair bonding in a double helix
confirmation, such as for double stranded DNA in a double-helical
confirmation.
"Confine region" refers to that portion of the nanopassage wherein
the double-stranded polynucleotide must deform to enter, traverse
and/or exit. In the examples provided herein, the deformation is
achieved by applying an electric field across the nanopassage such
that the polynucleotide, which is charged, experiences a force
sufficient to deform or stretch thereby permitting entry, traverse
and/or exit through the confine region and, therefore, traverse the
nanopassage. When the force on the polynucleotide is removed, such
as by lowering or removing the applied electric field, the
polynucleotide cannot enter, traverse and/or exit the confine
region. In an aspect only a portion of the nanopassage is a confine
region, such as a portion that is centrally located with respect to
the membrane surfaces. In another aspect, substantially all or all
of the nanopassage is the confine region.
Accordingly, "driving voltage" refers to the voltage applied across
the nanopassage required to initially force the polynucleotide into
the nanopassage, and specifically into the confine region of the
nanopassage. "Threshold voltage" refers to the voltage across the
nanopassage required, once the polynucleotide has entered the
confine region, for the polynucleotide to unidirectionally traverse
through the confine region and out of the nanopassage. "Holding
voltage" refers to a voltage that is sufficiently low that the
portion of the polynucleotide in the confine region is unable to
exit the confine region. In this manner, the holding voltage is
less than the threshold voltage and the driving voltage. In an
aspect, the threshold voltage is less than the driving voltage,
such that polynucleotide controllably traverses the nanopassage and
confine region in a manner that is electrically measurable.
Generally the holding voltage is greater than zero (e.g., the
polarity of the electric field is not reversed). However, so long
as the confined portion of the polynucleotide cannot move in the
opposite direction, the holding voltage can, if desired, be less
than zero (e.g., reversed polarity).
"Fluid communication" refers to a nanopassage that permits flow of
electrolyte, and specifically ions in the electrolyte from one side
of the membrane (e.g., first fluid compartment) to the other side
of the membrane (e.g., second fluid compartment), or vice
versa.
"Deforms" refers to the membrane nanopassage having a geometry that
makes it not possible for the polynucleotide to traverse or move in
a direction from one fluid compartment to the other without the
polynucleotide changing conformation. In particular, with respect
to the double helix geometry, it is necessary for the axial rise
between adjacent base pairs to increase before that portion of the
polynucleotide can enter the confine region. This increase in axial
rise is achieved by applying an electric field across the
nanopassage such that polynucleotide. This change in axial rise is
also referred to as stretch.
"Confine state" refers to whether a nucleotide or nucleotide base
pair is uniquely positioned in the confine region or whether a
nucleotide or nucleotide base pair is not uniquely positioned in
the confine region, such as a base pair that is moving out of the
confine region.
"Current blockade" refers to the measured current, at the holding
voltage when there is a nucleotide base or nucleotide base pair
trapped in the confine region. The magnitude of the current
blockade depends on the type of base or base pair in the confine
region including base identity and base methylation state. For
embodiments where there is more than one base or more than one base
pair in the confine region, the plurality of bases in the confine
region will affect the current blockade and sequential measurements
of current blockade as the polynucleotide traverses the confine
region may be used to uniquely identify bases and base pairs.
The methods provided herein are capable of characterizing double
stranded DNA such as by evaluating the percentages of base type,
providing sequence determination over a select length of the
polynucleotide, or sequencing entire lengths. In addition,
characterizing may refer to assessing a methylation parameter,
state and/or pattern.
The invention may be further understood by the following
non-limiting examples. All references cited herein are hereby
incorporated by reference to the extent not inconsistent with the
disclosure herewith. Although the description herein contains many
specificities, these should not be construed as limiting the scope
of the invention but as merely providing illustrations of some of
the presently preferred embodiments of the invention. For example,
thus the scope of the invention should be determined by the
appended claims and their equivalents, rather than by the examples
given.
Example 1
Synthetic Nanopore for Sequencing Double-Stranded DNA
Sequencing a single molecule of DNA extracts the maximum amount of
information from a minimum of material and pre-processing. When
paired with a high-throughput and low cost sequencing instrument,
it could change the flow of genomic data from a trickle to a
deluge--thrusting genomics within the grasp of personalized
medicine. Provided in this example is a nanopore instrument that
sequences the genome in a cost effective manner, currently for less
than $1000. Nanopore sequencing relies on the electric signal that
develops when a single, polyanionic DNA molecule is forced by an
electric field to translocate through a pore. The nanopore
sequencing concept is revolutionary because it combines the
potential for long read lengths (>5 kbp) with high speed (1
bp/10 ns), while obviating the need for costly and error-prone
procedures like PCR amplification due to the exquisite single
molecule sensitivity. However, high fidelity reads demand stringent
control over both the molecular configuration in the pore and the
translocation kinetics. The molecular configuration determines how
the ions passing through the pore come into contact with the
nucleotides, while the translocation kinetics affect the time
interval in which the same nucleotides are held in the constriction
as the data is acquired. Until now, no nanopore prototype proffered
for sequencing has shown any prospect of satisfying both of these
specifications at the same time.
A solid-state pore with a diameter (d) smaller than the effective
diameter of the polynucleotide, e.g., the double helix, is used to
sequence double-stranded DNA (dsDNA). We have shown that there
exists a voltage threshold for dsDNA permeation through pores
d<3 nm, consistent with the notion that the molecule must be
stretched by the applied electric field to translocate through the
pore..sup.2-4 We show here that if the voltage is rapidly switched
from a value above the stretching threshold to zero during a
translocation, dsDNA can be held in a pore indefinitely with a
base-pair (bp) trapped in the constriction, for sufficiently small
pore diameters (i.e. d<2 nm for double stranded DNA). Our recent
experiments with d.about.2.5 nm pores indicate that we can weakly
trap .lamda.-DNA for as long as 56 s, which is about 59,000.times.
longer than the duration of a typical translocation (900 .mu.s)
observed for voltages above the stretching threshold..sup.5
Moreover, stretching dsDNA causes the bps to tilt as they
translocate through the pore, which in turn modulates the
electrolytic current. Accordingly, we measure a sequence-dependent
blockade signal when the molecule is trapped, to discriminate A-T
from C-G bps. One concern with sequencing dsDNA in this manner
relates to determining which nucleotide is on which strand, e.g.
distinguishing A-T from T-A. However, our molecular dynamics (MD)
simulations show that the orientation of the by tilt caused by the
confinement is maintained during translocation with the nucleotides
on one strand always lagging their complement on the other. This
combined with a nanometer-scale slit geometry for the pore that
forces the current near the nucleotides indicates that a unique
sequence of dsDNA can be characterized by first trapping the
molecule and then using lock-in measurements of the blockade
current to read bps with 3 pA precision.
Using this strategy to sequence a genome involves satisfying two
requirements: 1. the ability to capture and trap a long dsDNA
molecule in a d.about.2.0 nm pore; and 2. a nanopore with
signal-to-noise performance commensurate with 3 pA current
resolution. Corresponding to these requirements, this example
specifically focuses on two items:
1. Determining conditions for trapping a long (>5 kbp) dsDNA
molecule in a nanopassage, such as a 2.0.times.1.0 nm nano-slit. A
long read length is a primary advantage associated with nanopore
sequencing, but our experience shows that it is difficult to
capture DNA >5 kb in <2.5 nm diameter pores. On the other
hand, we observe that short DNA strands (<100 bp) can be forced
to translocate through the pore at a high rate. Long DNA strands
are forced away from the high field region in the constriction by
the electro-osmotic flow through the pore. Thus, the capture rate
is adversely affected by the large radius of gyration, which
depends on the DNA persistence and contour lengths, and the pore
surface charge, and the large voltage required to stretch dsDNA. We
fabricate and test nanometer-scale 2.0.times.1 nm slits in
ultra-thin (.about.10 nm) silicon nitride membranes, formed by
electron-beam ablation in an electron microscope with a probe
corrector to improve the brightness of the smaller diameter
electron beams. Then, using pH to soften the DNA along with
chemical surface treatments and atomic layer deposition to modify
the pore charge, we further analyze conditions for trapping dsDNA,
informed by quantum/molecular mechanics (QM/MM) simulations.
2. We also produce a low noise, high sensitivity instrument having
a nanopassage suitable for sequencing dsDNA. To satisfy the
electric field specification for stretching, the signal-to-noise
required for single by discrimination, and the high-speed
requirement for stepping between bps in a trap, we can fabricate
nano-slits ranging from 2.5.times.1 nm to 1.6.times.0.7 nm in
ultra-thin (.about.10 nm) nitride membranes, using MD in
combination with Brownian dynamics to discover a pore geometry that
maximizes the blockade signal. To mitigate the noise we can
minimize the parasitic membrane capacitance associated with the
substrate by using 1 .mu.m.times.1 .mu.m nitride membranes 10 nm
thick fabricated on a sapphire substrate. Together these efforts
should produce a nano-slit instrument with a bandwidth >330 kHz
and rms-noise specification <0.4 pA suitable for reading a bp,
that can step at high speed from one by to the next for
re-sequencing a genome for less than $1000.
To sequence DNA using a nanopore, one must first find a robust,
nanoporous structure of an appropriate size--comparable to or even
less than the size of DNA to maximize the signal. The prospects for
low cost, high-throughput nanopore sequencing are currently being
explored using as prototypes either .alpha.-hemolysin (.alpha.-HL)
and its mutants, or nanopores in solid-state membranes. Since the
translocation velocity through the pore can be very high--about 1
bp/10 .mu.s for .alpha.-HL and .about.1 bp/10 ns for a solid-state
pore--a single DNA molecule may be sequenced quickly and
inexpensively, so long as the bases are discriminated electrically.
Single base resolution on a translocating strand has not been
demonstrated in the art. High fidelity reads demand stringent
control over both the molecular configuration in the pore and the
translocation kinetics. Control of the molecule configuration
determines how the ions passing through the pore come into contact
with the nucleotides in the constriction, while the translocation
kinetics affect the time interval in which the same nucleotides are
held in the constriction and data is acquired. Until now, none of
the nanopore prototypes proffered for sequencing have shown any
prospect of simultaneously satisfying both of these
specifications.
Kasianowicz et al were among the first to adopt .alpha.-HL
nanopores to detect and sort single DNA molecules.20,21 .alpha.-HL
is a mushroom shaped heptamer that assembles across a phospholipid
membrane. It is composed of seven identical subunits arranged
around a central axis; the transmembrane portion is a .beta.-barrel
about 5 nm long with a minimum diameter of 1.5 nm. By placing this
protein within a lipid membrane, an electric field induced flow of
ions through the protein can be measured. If ssDNA or RNA is added
to the anodic side, the translocation through the .alpha.-HL pore
and resulting current blockade can be detected. The correspondence
between current blockades and translocation of DNA between
compartments was demonstrated by quantifying the DNA in the
cathodic compartment using competitive PCR.20
There are limitations to using .alpha.-HL for sequencing, however.
First are the obvious structural limitations--the protein structure
is difficult to change in a predictable way. Though it is possible
to introduce subtle mutations into the protein, gross structural
changes are inordinately difficult. Chief among these structural
limitations in .alpha.-HL are the length of the nanopore, and hence
the thickness of the membrane, and the diameter of the pore. For
example, the .alpha.-HL channel is only 1.5 nm in diameter--it will
not admit dsDNA. The shape and length of the nanopore also means
that it is functionally impossible to measure only one base at a
time, making the sequence, which lies within the pore nontrivial to
interpret, as multiple nucleotides are contributing to the signal.
Finally, the lipid bilayer presents another limitation. The lipid
bilayer membrane, which is usually suspended over a Teflon orifice,
is typically 25-100 .mu.m in diameter and only 5 nm thick. It
ruptures after a few hours of use or after cycling the electrolyte
a few times and the large size of the membrane produces a
capacitance that adversely affects the frequency and noise
performance.
Bayley et al.22 recently engineered .alpha.-HL in such a way to
improve the signal to noise ratio, i.e. to hold a nucleotide in
place for a longer period of time, in order to perform more
averaging. By modifying the .alpha.-HL such that a cyclodextrin is
placed in the .beta.-barrel, the time that the pore is occluded by
a single nucleotide can be extended. This allows more accurate
measurements (>90%) of what nucleotide is in the pore based on
blockade current. This method was used in combination with an
exonuclease to determine the composition of ssDNA using an
exonuclease to cleave off individual dNMPs, and then measure them
as they are captured by the .alpha.-HL nanopore. This could
potentially be used on raw, genomic dsDNA, and is even sensitive to
base modifications such as cytosine methylation. However, it
suffers from a crippling problem of logistics: i.e. how to
transport the cleaved nucleotides from the exonuclease to the pore,
ensuring that they arrive in the same sequence as found in the
original DNA strand, that none escape (missing a base), and that
the exonuclease does not outpace the pore. Tethering the
exonuclease to the nanopore has been proffered as a solution, but
this scheme is a nontrivial extension to the original nanopore
sequencing concept.
We use a solid-state pore with a diameter smaller than the double
helix to sequence a dsDNA. In contrast to .alpha.-HL, the size and
shape of the pore in a solid-state membrane can be controlled on a
sub-nanometer scale, allowing for a specific geometry to be
tailored to the purpose. Solid-state pores also offer vastly
improved stability: they are resilient in much harsher chemical and
thermal environments useful for denaturing the DNA, as well as
allowing for easier integration with other electrical or
microfluidic components. Through microfabrication techniques, the
solid-state membrane can be reduced to sub-micrometer scales, in
principle, mitigating parasitic capacitance effects and improving
electrical performance. There are several different methods
available to create nanopores in thin membranes, such as ion-beam
milling,23 ion-track etching,24 silicon dioxide reflow25 or
electron-beam ablation.26 These techniques may be used on a variety
of different membrane materials--allowing for different chemical
properties, such as surface charge density, and electrical
properties, such as capacitance. Thus, semiconductor
nanofabrication technology is a key aspect of our approach.
To sequence dsDNA we first trap a molecule in a pore with d.about.2
nm; and then use low-noise, lock-in measurements of the
sequence-dependent blockade current to discriminate between bases.
There exists a voltage threshold for permeation of dsDNA through
pores <3 nm in diameter, consistent with the notion that the
molecule must be stretched by the applied electric field to
translocate through the pore.2-4 We show that if the voltage is
rapidly switched during a translocation from a value of above the
stretching threshold to value below, it is possible to weakly trap
.lamda.-DNA for as long as 56 s in a d.about.2.5 nm pore, which is
about 59,000.times. longer than the duration of a typical
translocation (900 .mu.s) observed for voltages above the
stretching threshold. Moreover, stretching dsDNA causes the
base-pairs to tilt as they translocate through the pore, which in
turn modulates the electrolytic current. Accordingly, we measure a
sequence-dependent blockade signal when the molecule is trapped,
suggesting we can discriminate A-T from C-G bps. Thus, our findings
indicate that nanopores with d.about.2.5 nm afford us some control
over both the translocation kinetics and the molecular
configuration in the pore. Performance may be optimized and
improved, making it suitable for high-throughput low cost
sequencing.
In the sequencing process, we operate a nanopore like tweezers,
repeatedly trapping dsDNA in the pore while performing high
frequency, narrow band, lock-in measurements to resolve signatures
of the base-pairs in the pore current. FIG. 1 illustrates
schematically an exemplary process for sequencing DNA. The
experimental apparatus has two chambers, each filled with
electrolyte separated by a membrane with a nanopore in it that is
<3 nm in diameter. dsDNA is injected in the cis chamber. A
driving bias (see FIG. 1(h)), V0, is then applied and a
corresponding ionic current (FIG. 1(i)), I0, flows in the absence
of dsDNA through a nanopore (FIG. 1(a)). The driving bias causes
dsDNA in the vicinity to migrate towards the pore and eventually it
is captured by the field as shown in FIG. 1(b). The driving bias
above threshold stretches the DNA, enabling it to thread the pore
as shown in FIG. 1(c). At the onset of a current blockade, when a
sequence-dependent blockade current, I1, is detected, feedback is
used to reduce the voltage to V.sub.1 and trap the molecule. At
this time a lock-in measurement of the current, I2, is used to
discriminate between the bases. After the measurement, the DNA is
induced to advance by applying a voltage pulse greater than the
threshold for the translocation--such as by moving one base at a
time through the pore if switched sufficiently fast--until the
current, I3, indicates the location of another base. And then the
cycle continues from a measurement of I3 to a measurement of
I4.
Requirements of a nanopore instrument for sequencing dsDNA include:
1. The geometry of the constriction must be stringently controlled
as it determines the sensitivity and the electric field
distribution. 2. The frequency response and noise performance have
to be commensurate with switching the membrane voltage on a 10 nsec
time scale for stepping from one base-pair to the next and
detecting a single base-pair. 3 The molecule must be trapped in the
pore for sufficient time to discriminate individual base-pairs.
These three aspects are addressed in this example.
Fabrication of a Solid-State Nanopore <2.6 nm in Diameter. A
unique aspect of the approach provided herein relates to the pore
geometry; the thickness and composition of the membrane can all be
controlled with sub-nanometer precision using semiconductor
nanofabrication practices. This precision translates directly into
control of the distribution of the electric field, (which has
already led to the development of the most sensitive device for
charge measurement: the single electron transistor.) Following
several innovations23-26 in the fabrication of solid-state
nanopores, we develop methods for producing nanometer diameter
pores in robust membranes as illustrated in FIG. 2. For example, we
produce silicon nitride membranes by depositing an LPCVD Si3N4
film, ranging from 30 nm to 200 nm thick (nominally), on the top of
a 300 .mu.m thick (float-zone) Si handle wafer. To control the
hydrophobicity, we control the amount of oxygen, silicon and
nitride in the film. To reduce the thickness, either the nitride
membrane is sputtered in a 5 .mu.m.times.5 .mu.m area using
focused-ion beam milling or it is uniformly etched in 20:1 H2O:49%
HF for 30-40 min at room temperature. Afterward, a polyimide
photoresist with thickness of 3.6.+-.0.6 .mu.m is spin deposited on
top of the chip, and a 5 .mu.m window is opened over the membrane
using UV lithography as shown in FIG. 2(a). The polyimide is used
primarily to reduce the parasitic substrate capacitance.
A nanometer-size pore is subsequently sputtered into membranes like
these using a tightly focused (1.6 nm spot-size) 9.degree. .alpha.
(cone angle), high energy (200 kV) electron beam emanating from a
JEM-2010F transmission electron microscope (TEM) operating in
convergent beam diffraction mode. Using TEM images taken at
different tilt angles, we model the pore geometry as two
intersecting cones (bi-conical) each with >20.degree. cone
angle.26 By stringently controlling the beam conditions and
membrane thickness (guaranteed by Electron Energy Loss
Spectroscopy) it is possible to produce pores with practically the
same geometry with sub-nanometer precision, as illustrated by the
array in FIG. 2(d).
We use a new tool, a JEOL-2200F, which is an upgraded version of
the JEM-2010F. The 2200F has a piezo-stage for atomically precise
control over the sample position, and an aberration probe corrector
that allows for increased (8.times.) brightness with a smaller
probe. This corrector enables us to sputter with a smaller (<1.6
nm) spot which, in combination with the piezo-stage, provides more
precise control over the pore geometry. This system facilitates the
production of various nanometer-sized passages, such as a
nanopassage having a 2.0.times.1.0 nm nano-slit with a 1 nm beam to
closely conform to the twisting, propeller-like, helical shape of
B-form dsDNA.
Electrical Characterization of a Solid-State Nanopore.
B-form dsDNA is a stiff, highly charged polymer with a solvated,
helical structure about 2.6-2.9 nm in diameter, according to
neutron scattering, that depends on the sequence and the number of
strongly bound water molecules included in the primary hydration
shell. So, precise, sub-nanometer control over the geometry of the
nanopore, and the thickness and composition of the membrane
translates directly into control of the distribution of the
electric field, and accordingly the configuration of dsDNA in the
constriction during a translocation.
When an electric field is applied across a membrane with a
bi-conical pore d<3 nm in it that is immersed in electrolyte,
the voltage is effectively focused onto that portion of the
molecule near the center of the membrane over a region about 1-3 nm
wide.2,3 This means that dsDNA has to first diffuse within range of
the pore to be driven through it by the electric field. The rate of
DNA capture is roughly given by R=2.pi.CDr, with R the capture
rate, C the concentration of DNA, D the diffusion constant of DNA
in free solution, and r the radius of probable capture by the pore,
dependent on the voltage applied.17 Once it is inside the pore,
there are three main forces that affect the DNA. The first and
strongest force is the electric field, acting primarily on the
negatively charged phosphate backbone of DNA. The electric field
causes electrophoretic motion of the DNA, driving it forward into
the pore while the positively charged ion cloud surrounding it is
driven back. There is an electrostatic interaction with the pore
walls, and/or a nonpolar (van der Waals) interaction. And finally,
there is a drag force associated with the movement of the polymer
in solution, which is essentially a frictional force.
To determine the microscopic origin of the net force exerted on DNA
in a nanopore at a given transmembrane bias, we use MD to simulate
the system.27 We find three regimes for the dependence of the net
force F on the applied electric field E, which we categorize
according to the pore diameter. For a pore diameter >5 nm, the
interactions with the pore itself are negligible, which makes sense
considering the small Debye length (.about.1 nm) and the weak
interaction of the van der Waals (r-6 dependence). When the pore
diameter is between 3.6 and 5 nm, the electrolyte still behaves as
it does in bulk solution, but direct interaction between the DNA
and the pore surface becomes important. Finally, when d<3.6 nm,
the viscosity of water in a thin film between DNA and a nanopore
surface is larger than in the bulk and depends on the shearing
velocity if DNA is moving. In this regime, the interactions between
DNA and the pore can be much stronger and the microscopic details
of the pore surface strongly affect the friction. A nonlinear
dependence of the force on the applied electric field is expected,
which is optimal for sequencing, as it allows the force and
velocity of DNA translocation to be easily affected. Moreover, it
forces DNA to move into and through the pore single file as more
than one double helix cannot fit in the pore at the same time,
occluding the electrolytic current through the pore and maximizing
the signal.
When forced into a pore smaller than the double helix, the leading
edge of the dsDNA penetrates the membrane to a constriction
.about.2.5 nm in diameter. If the differential force acting on the
leading nucleotides is insufficient to stretch the helix, the
translocation stalls there, but as the bias increases and the
differential force exceeds that required to stretch dsDNA, the
molecule is pulled towards the center of and eventually through the
membrane. The two DNA strands do not pass through pores with
diameters 1.6<d<2.5 nm in the same way, however. The
confinement causes the basepairs to tilt.2,3 For diameters <1.6
nm, dsDNA unzips and the strands translocate through the pore one
at a time.3
We test the electromechanics of dsDNA in a nanopore with a
2.5.times.2.0.+-.0.2 nm cross-section--smaller than the DNA double
helix--in a membrane 15.0.+-.2.2 nm thick shown in FIG. 3(a). (This
represents an early attempt to fabricate a nano-slit using the
2010F. Although the pore is elliptical, the beam diameter
.about.1.6 nm is too large to effectively sputter a slit with
eccentricity .about.1.) When .lamda.-DNA is injected into the
electrolyte at the negative (cis) electrode and 800 mV applied
across the membrane, current transients like those shown in FIG.
3(b) are observed. The current transients occur randomly as a
function of time as illustrated in the figure, but the interarrival
time decreases with increasing concentration of DNA on the cis side
of the bi-cell. There is a variety of transients, some of which are
illustrated in FIG. 3(c). The distribution of transients can be
represented by a blockade with a peak value at 62.+-.11% of the
open pore current (i.e. .DELTA.I/I0=0.62.+-.0.11) as shown in FIG.
3(d). A blockade is supposed to be the reduction of the
electrolytic current through the pore due to the translocation of
DNA. In support of this interpretation, MD estimates indicate a
.DELTA.I/I0=0.4.+-.0.1 blockade through a 2.0 nm diameter pore in
100 mM KCl.
FIG. 3(b) illustrates threshold behavior, showing a dearth of
transients found in a current trace measured at 200 mV in
comparison with 800 mV. FIG. 3(e) summarizes the frequency of
blockade events observed as a function of the voltage applied
across the membrane over the range from 100 mV to 1V. Generally, we
observe that the number of blockades rises abruptly over a range of
.about.200 mV near a threshold that is sensitive to the pore
diameter. If we assume that each blockade corresponds to dsDNA
permeating the pore, then the permeation rate can be described by
the transition-state relation of the Kramers type:4
R=R0V/(1+exp[q*(U-V)/kT], where R0 is a frequency factor, q*U is
the effective barrier height, q*V is the reduction in the energy
barrier due to the applied potential, and kT is the thermal energy.
Using this relation, the data are fit and the results overlaid on
the scatter plot in FIG. 3(e). We find a threshold of
U=0.46.+-.0.02V with q*=0.8.+-.0.2e, which presumably corresponds
to the force required to stretch the leading nucleotides in the
pore--below this voltage DNA is not supposed to permeate the
membrane. This conclusion is supported by prior work2-4 which shows
that the threshold depends on pH, pore diameter, membrane
thickness, dsDNA sequence and methylation profile. Typically, for a
2 nm diameter pore in a 10 nm thick membrane at pH 8, the threshold
voltage is .about.3V.
The duration of the blockade grows longer with the DNA contour
length,26 which supports the interpretation of the blockade current
as a translocation across the membrane. The dependence of the
blockade duration on the membrane voltage offers more support for
this interpretation. FIG. 3(f) shows the frequency of current
transients associated with .lamda.-DNA as a function of duration
with the voltage as a parameter. If the blockade duration
corresponds with the interval that DNA blocks the pore, then the
average transient width tD signifies the time required for 48.502
kbp .lamda.-DNA to translocate through the pore. We find that
tD=0.160.+-.0.01 ms, 0.53.+-.0.06 ms, 1.1.+-.0.1 ms, 0.82.+-.0.07
ms and 2.49.+-.0.25 ms for voltages of 1.0V, 800 mV, 700 mV, 600 mV
and 500 mV respectively. The corresponding translocation velocity
estimated from the quotient ranges from 1 bp/3.3 ns at 1V or 1
bp/11 ns at 800 mV to 1 bp/50 ns at 500 mV, which is consistent
with MD simulations performed under similar conditions. The inset
to FIG. 3(f) shows a plot of the voltage dependence of the
reciprocal of the average transient width, i.e. 1/tD, measured
above threshold. 1/tD falls abruptly near the threshold value,
which is consistent with the idea that the molecule may become
trapped in the pore near the threshold voltage. The line in the
inset is a least-squares fit to the data, which has a slope of 11
V.sup.-1 s.sup.-1 with a voltage-intercept of 0.53V, which is
comparable to the threshold voltage inferred from FIG. 3(e).
These observations are in sharp contrast with prior work on larger
diameter pores,29,30 and measurements that we performed on a
3.6.times.3.2.+-.0.2 nm pore with a cross-section larger than the
DNA double helix cross-section in a 31.5.+-.2.0 nm thick nitride
membrane. We observe current blockades at much lower transmembrane
bias in the 3.6.times.3.2 nm pore--into the millivolt range. If we
assume that each blockade corresponds to dsDNA permeating the pore
and fit to the transition-state relation, we find a threshold
voltage of U<0.06.+-.0.02V with q*=1.0.+-.0.2e (data not shown.)
And if the blockade duration corresponds with the interval that DNA
blocks the pore, and the average transient width tD signifies the
time required for 48.502 kbp .lamda.-DNA to translocates through
the pore, then for the 3.6.times.3.2 nm pore, tD=0.031.+-.0.007 ms
0.0321.+-.0.007 ms, 0.0677.+-.0.003 ms, and 0.403.+-.0.26 ms for
voltages of 600 mV, 400 mV, 200 mV and 100 mV, respectively, which
is consistent with prior estimates of the translocation velocity
>1 bp/10 ns.29,30
The high translocation velocity observed in a solid-state nanopore
at the high voltage required to stretch DNA demands a nanopore with
a high frequency response to detect blockades, otherwise they
cannot be resolved reliably (and so typically we use quantitative
PCR to count the molecules that permeate the membrane through the
pore.) FIG. 3(g) shows the frequency responses of four types of
membranes: two associated with different nitride thicknesses, 12 nm
and 200 nm, on a silicon substrate, a third associated with a
composite 30 nm nitride membrane coated with a 3.6 .mu.m polymer
film, all on a silicon substrate, and a fourth a simulation of a 12
nm nitride membrane on a sapphire substrate--each membrane contains
a nanopore ranging in diameter from 2-5 nm. Generally, we find that
the current frequency response consists of two components: one
associated with the conductance through the pore, which
predominates at low frequency and is manifested by zero-slope
versus frequency; and another due to the displacement current
associated with the membrane capacitance and associated parasitics.
While both depend linearly on the applied voltage, the displacement
current increases with frequency, which is why the current grows so
large at high frequency.
Detailed models that precisely capture the frequency response of
the nanopores (FIG. 3(h)), which are rooted in the physical
structure and reflect, in a limited way, the distributed nature of
the electrical parameters are developed. In addition to the
capacitance of dielectric materials such as polyimide, Si3N4, the
models also account for the depletion layer capacitance in the Si
handle, the dielectric loss in each case, the resistivity of the
substrate, the resistivity of the KCl electrolyte, the double layer
that is associated with the interface between a charged surface and
an electrolyte solution, and the Faradic impedances associated with
charge transfer: Fits to the data are represented by the solid
lines in FIG. 3(g): the models accurately account for the frequency
response. For economy, to illuminate the relationship between the
ac pore current, i, and the ac voltage, v.sub.m, responses across
the membrane, we use the simplified model shown in FIG. 3(i) due to
Smeets et al.31 This model is comprised of an effective capacitance
Cm representing the membrane in parallel with the pore resistance
Rp and in series with the electrolyte/electrode resistance Rel.
According to the model, the Fourier transform of the frequency
response functions are given by:
I.function..omega..omega..times..times..times..function..omega..times..ti-
mes..times..times..upsilon..fwdarw..upsilon..function..omega.I.function..o-
mega..times..omega..times..times..times..function..omega..times..times..ti-
mes..times..upsilon..apprxeq..omega..times..times..times..times..upsilon.
##EQU00001##
so that v.sub.m(t)=v.sub.ine.sup.-t/CmRel. Thus, the transient
voltage response time is determined by the time constant: Cm Rel.
If Cm is reduced by mitigating the effect of parasitics, then the
high-frequency response is determined by the time constant
associated with the window capacitance and the load resistance,
which consists of the amplifier load in series with the
electrolytic resistance. The window capacitance is controlled by
the membrane thickness and the area, while the series resistance
can be minimized by increasing the electrolyte concentration. For
example, FIG. 3(g) illustrates the dramatic effect that parasitics
associated with the substrate have on the frequency response. After
replacing the handle wafer with sapphire instead of silicon, the
pore conductance now predominates the current response up to a
frequency of fz0=1/2.pi.RpCm.about.200 Hz and the membrane
capacitance is substantially reduced to Cm=1.6 pF. With this
capacitance and 1M KCl, the voltage response time is .about.4 nsec,
which is fast enough to move from one base-pair to the next at 1
bp/10 ns.
In addition to the capacitance of dielectric materials such as
polyimide and Si3N4, the models also account for the depletion
layer capacitance in the Si handle, the dielectric loss in each
case, the resistivity of the substrate and the KCl electrolyte, the
double layer that is associated with the interface between a
charged surface and an electrolyte, and the Faradic impedances
associated with charge transfer. To determine the parameters
governing the model, which are delineated in Table 1, the values of
the various lumped elements were first estimated from the geometry
and then the data was fit using a least-squares minimization
algorithm to converge to the final values. Fits to the
corresponding models are represented by the solid lines in FIG.
3(g).
This model accurately accounts for the frequency response of the
current. And since it is derived from the physical structure, it
can be used to elucidate strategies for improving the
signal-to-noise through changes in the pore-membrane structure. For
example, the membrane voltage determines the electric field in the
pore, which affects the translocation kinetics as well as the
potential barrier associated with the molecule in the constriction
and therefore the blockade current. To illuminate the relationship
between the ac pore current, i, and the ac membrane voltage,
v.sub.m, response across the membrane, we used for economy the
simplified version of the model of FIG. 3(h) shown in FIG. 3(i).
This model is comprised of an effective capacitance Cm representing
the membrane in parallel with the pore resistance Rp and in series
with the electrolyte/electrode resistance Rel. According to the
model, the Fourier transform of the frequency response functions
and the corresponding membrane voltage response are provided so
that v.sub.m(t) is obtained, as described above. Thus, the
transient voltage response time is determined essentially by the
time constant: CmRel. As illustrated in Table 1, typically, fits to
the data yield effectively Cm=100 pF and Rel=10 k.OMEGA. for 1M KCl
electrolyte so that .tau..about.1 .mu.sec. If Cm or Rel is reduced
by mitigating the effect of parasitics, then the high-frequency
response can be improved.
The so-called "membrane" capacitance is actually dominated by
parasitic capacitances associated with the seal area over the
handle wafer and the cabling, while the electrolyte resistance is
determined mainly by the electrolyte concentration and the Ag/AgCl
electrodes geometry relative to the pore. We can reduce the
parasitic capacitance by using: 1. thicker Si3N4 membranes; 2.
composite membranes consisting of thick polyimide on a thin
miniaturized Si3N4 membrane; or by 3. eliminating depletion in the
handle wafer; or 4. eliminating cabling; and/or by using
capacitance compensation through external circuitry, which has been
used successfully for patch clamping. While capacitance
compensation can provide a vast improvement in the high frequency
performance, it also contributes noise,5 so mitigation of the
parasitic capacitance through miniaturization of the membrane still
offers the most promising route to high fidelity electrical
measurements as evident from the improvement illustrated by 30 nm
thick nitride/polymer composite membrane shown in FIG. 3(a). FIG.
3(a) also illustrates the effect the parasitic capacitance
associated with the substrate has on the frequency response. After
replacing the handle wafer with sapphire instead of silicon, the
pore conductance now predominates the current response up to a
frequency of fz0=1/2.pi.RpCm.about.200 Hz and the membrane
capacitance is substantially reduced to Cm=1.6 pF. With this
capacitance and 1M KCl, the membrane voltage response time becomes
CmRel.about.4 nsec, which is faster than the DNA translocation
velocity (1 bp/10 ns).
Further improvements can be gleaned from miniaturization and
repositioning the Ag/AgCl electrodes closer to the pore. For a
micro-disk geometry embedded between two dielectrics, it can be
shown that the resistance follows the law: (1/4R)(1/t), where R is
the radius and t the exposed thickness. Thus, we can use a
composite nitride/polyimide membrane 1 .mu.m.times.1 .mu.m in area
with embedded Ag/AgCl electrodes 1 .mu.m thick sandwiched between
the polyimide and the nitride, 1 .mu.m in diameter encircling the
membrane, all on a sapphire substrate. We estimate electrolyte
resistance to be .about.500 for 1M KCl--representing an improvement
of 200.times..
We expect the largest signal for the smallest pore diameter, but if
the bandwidth is too narrow, it is difficult to resolve the
signature associated with the translocation of a single base-pair.
(Repeated measurements made using multiple pores with multiple
copies of DNA or multiple passes with a single molecule doesn't
really solve this problem.) So, following Smeets et al.31 we sought
a compromise between signal-to-noise and bandwidth by analyzing in
detail the electrical characteristics of solid-state nanopores,
representing them by the equivalent lumped element circuit shown in
FIG. 3(i).
The noise power spectra, corresponding to the measured frequency
response of the same three nanopores in FIG. 3(g), are shown on
FIG. 4(a) along with the spectrum of a 300 M.OMEGA. resistor, a
value comparable to the pore resistance. We analyze the noise into
components that can be categorized as: 1. thermal noise associated
with the resistance of the electrolyte and the pore resistance 2.
1/f or flicker noise that is related to the carrier density in the
pore; 3. dielectric noise associated with the membrane and holder;
and 4. noise originating with the measurement amplifier. FIG. 4(a)
shows the measured noise spectra superimposed on the fit of the
thermal (black), 1/f (red), dielectric noise (cyan) and amplifier
noise (green) contributions along with the total (orange). The
thermal noise density associated with pore resistance, is
negligible over the band. At low frequencies, the noise power
density is inversely proportional to the frequency, which is
indicative of the presence of excess, or 1/f, noise. Its noise
power spectrum is modeled by:
S.sub.1/f=I.sup.2.alpha./N.sub.cf.sup..beta., where I is the
current through the device, .alpha. is the Hooge parameter (an
empirically determined proportionality constant), Nc is the total
number of current carriers, f is the frequency, and .beta. is an
exponent that is typically unity. This portion of the spectrum can
be described with .beta.=1.085.+-.0.010, depending on the
electrolyte concentration. At frequencies >1 kHz, 1/f noise
becomes negligible, and the spectrum exhibits linear frequency
dependence up to about 50 kHz. The noise in the range 1-50 kHz is
dominated by dielectric noise with a spectrum of the form:
S.sub.d=4k.sub.BTDC.sub.D(2.pi.f), where kB is Boltzmann's
constant, T is the absolute temperature, and D and CD are the loss
tangent and capacitance of the dielectric material. This
capacitance and loss tangent are directly related to the lumped
elements comprising the circuits in FIG. 3(i): e.g. the loss
tangent is the tangent of the angle between the Cm capacitor
impedance vector and the negative reactance. Finally, above 50 kHz,
the spectrum is strongly affected by the bandwidth of the amplifier
(.about.55 kHz). The amplifier noise (referenced to the amplifier
input) has a density spectrum:
S.sub.amp=4.pi..sup.2f.sup.2.di-elect
cons..sub.s.sup.2Cm.sup.2/(1+4.pi..sup.2f.sup.2.tau..sub.sr.sup.2)
where .di-elect cons..sub.s.sup.2 is the thermal voltage noise of
the series (KCl and AgCl electrode) resistance, .tau..sub.sr=RsrCm
and Rsr is the uncompensated series resistance. The dip >50 kHz
is due to the amplifier, which we have modeled as a low pass
filter.
From this analysis we deduce that the dielectric capacitance
dominates the total noise. FIGS. 4(b,c) shows how the rms-current
noise increases with bandwidth and capacitance over the range from
8 pF-900 pF. Thus, reducing the membrane capacitance is the key to
improving both the frequency and noise performance.
Trapping a Single DNA Molecule in a Nanopore:
The data of FIG. 3(f) indicates that dsDNA can be trapped in a
nanopore that is smaller in diameter than a canonical double helix,
if the membrane voltage is switched to a value below threshold
while the molecule is translocating through the pore. To examine
this further, first we force dsDNA into a 2.5 nm pore and then,
once a blockade in the current is detected, reduce the
transmembrane bias at high-speed while the molecule is still in the
pore. Once the dsDNA is in the pore, if the bias is reduced below
the stretching threshold, the pore functions like a trap in
resisting the motion of the molecule. FIGS. 5(a) and (b) show data
demonstrating that it is possible to weakly trap a single
.lamda.-DNA molecule once it is translocating through the pore by
switching the electric field at high-speed. During normal
operation, a transmembrane bias of 800 mV, which is above threshold
according to FIG. 3(d), is applied across the membrane resulting in
an open pore current >3 nA. Once the onset of a blockade like
that shown in the inset to FIG. 5(a) is detected by a
differentiator, a programmed delay of about 200 .mu.s is introduced
before a latch switches the voltage from 800 mV to 200 mV--a value
well below the threshold. All the while, the pore current is
monitored. According to the ac models we have developed, the
membrane voltage tracks the voltage applied to the Ag/AgCl
electrodes, but with a longer time constant (<500 ns).
Corresponding to the abrupt change in voltage, a current transient
occurs on a sub-microsecond time scale. The transient sometimes
saturates the current amplifier immediately after switching from
800 mV for an interval of about 1-3 ms, even with compensation.
(This occurs because i=CdV/dt is still too large since
dV/d/=0.6V/10 ns). Eventually, the current returns to the open pore
value, 10, but not before we observe a sharp transient like that
shown near t=7.709 s in FIG. 5(a).
We assume that transients like these are indicative of the
.lamda.-DNA molecule exiting the pore after 6.73 s. So, we reasoned
that the current blockade observed during the time interval from
t=0.9814 to 7.709 s must be evidence of a weakly trapped
.lamda.-DNA molecule in the pore. Consistent with this inference,
we do not observe a current blockade at 200 mV when there is no
onset of a blockade at 800 mV. Moreover, the 3.6.times.3.2 nm pore
at a constant bias of 200 mV shows a distribution of blockade
durations that peaks near 200 .mu.s, but does not exceed 1 ms,
which makes the long (6.73 s) duration shown in FIG. 5(a)
extraordinary. The fluctuations illustrated in FIG. 5(b) also
support the mechanism of pore current blocked by a .lamda.-DNA: we
observe that for t<7.709 s the amplitude of the current
fluctuations increases relative to the open pore value found for
t>7.709 s. FIG. 5(c) delineates the relative change of the
current fluctuations during the blockade from the open pore
current. Focusing on the data filtered with a low-pass 1 kHz
filter, the .DELTA.I/I0 width of the histogram taken from the
blockaded current is 0.28I0, while the open pore histogram measured
over the same 3 s interval is only 0.17I0 wide, indicating a signal
beyond the noise for t<7.709 s, that is likely due to base-pairs
translocating through the pore. Different base-pairs can be
resolved if the signal is averaged over a sufficient time.
If the molecule is weakly trapped in the pore, then the average
translocation velocity must have slowed substantially to a value of
about (48.5 kbp-200 .mu.s.times.1 bp/11 ns)/500 ms=1 bp/221 .mu.s,
which is about .times.20,000 times slower than the velocity
estimate for tD obtained at 800 mV. After repeating this type of
measurement hundreds of times on the same pore, a comparison
between the distribution of the duration of blockades observed with
a transmembrane bias of 800 mV, and the time that expires between
the triggered voltage switch from 800 mV to 200 mV and the return
of the current to the open pore value reveals the dichotomy
illustrated in FIG. 5(d). While the peak in the distribution
obtained at a constant bias of 800 mV occurs near tD=200 .mu.s, the
distribution found when the voltage is switched from 800 mV to 200
mV occurs near tD.about.500-600 ms. If we assume that the molecule
is weakly trapped, then the translocation velocity at the peak must
have slowed substantially to a value of about (48.5 kbp-200
.mu.s.times.1 bp/11 ns)/500 ms=1 bp/17 .mu.s, which is more than
.times.1500 times slower than the velocity at 800 mV. We do not
observe blockades >4 ms at constant bias, however, we have
observed blockades after switching as long as 6.7 s. The
distribution of the normalized blockade current .DELTA.I/I0 with a
constant bias of 800 mV is .DELTA.I/I0=0.62.+-.0.11 wide, while the
blockades measure after switching the voltage from 800 mV to 200 mV
is .DELTA.I/I0=0.46.+-.0.07 wide, which overlap within the
error.
In support of these conclusions, we have made similar findings on
three pores with similar geometries. FIG. 6(a) shows an interesting
event obtained from a 2.5 nm pore, which represents another trapped
.lamda.-DNA molecule. First, a bias of 600 mV (above threshold) is
applied across the membrane, and then, once the onset of a blockade
is detected, a delay of about 200 .mu.sec is introduced before the
voltage is switched from 600 mV to 100 mV (below threshold), while
the current is monitored. Eventually, the current returns to the
open pore value, near t=58.8 s in FIG. 6(a). We assume that this
transient is indicative of the molecule exiting the pore after 56
s, corresponding to a translocation velocity of >1 bp/1.8
ms.
With the molecule trapped under the conditions (i.e. for t<58.8
s), we filter the current data shown in FIG. 6(a) using a 20-1 kHz
bandpass filter and formed histograms of the current fluctuations
using 0.5 s windows. Each window shows a histogram similar to that
shown in FIG. 6(b), which can be represented by the superposition
of two Gaussian distributions: one (solid blue) offset from the
median (.DELTA.I=0) by .DELTA.I=+2.86 pA with a width of 7.6 pA;
and another (solid red) offset by .DELTA.I=-6.51 pA with a width of
5.8 pA. We attribute these separate peaks to resolved C-G/G-C and
A-T/T-A bps, respectively. The base-pairs can be resolved in this
case because of the correspondingly longer (.about.2 ms) time that
the base-pair is trapped in the constriction compared with the (200
.mu.s) trap time for the molecule in FIG. 5(a). In particular, as
shown in FIG. 6(d), the trapping or holding time (indicated as
dwell time on the x-axis), significantly increases when the voltage
is decreased from a threshold or driving voltage (600 mV)
sufficient to stretch the DNA to a correspondingly lower holding
voltage of 150 mV or 100 mV. The blockade duration increases from
about 900 .mu.s to about 200 ms.
Trapping a Single DNA molecule in a Nanopore (Simulation): We use
all-atom MD simulations in conjunction with Brownian Dynamics and
QM/MM methods to both visualize and test the interactions between
the dsDNA and the pore, and optimize the instrument for sequencing.
For example, MD simulations of the experiments described above
demonstrate that the motion of the dsDNA can be slowed or
effectively stopped when the driving voltage is turned off, that
there exists a threshold voltage for restarting the translocation
process and that the base-pairs can be resolved if signal is
averaged long enough. FIG. 7(a) illustrates the simulated system at
a 250 mV transmembrane bias, which includes an effectively infinite
fragment of dsDNA, a nanopore of a 2.6 nm.times.2.1 nm
cross-section, and 100 mM KCl. The molecular conformation in the
constriction is stretched beyond 0.34 nm per basepair by about
8-20%. DNA transport is observed only when a bias of 1V is applied,
as illustrated in FIG. 7(b). At 0.25 and 0.5V, the DNA's motion is
arrested, following a small initial displacement caused by
stretching. (This threshold should be smaller than the threshold
for stretching the leading nucleotides, since the molecule is
already in the pore.) By analyzing dsDNA's displacements at 0V, we
determine that the pore acts as a harmonic trap with an effective
spring constant of 3.0.+-.0.8 nN/nm, as shown in the FIG. 7(b)
inset.
Theoretical studies have shown that the probability of an escape
from a trap depends sharply on the force applied to the molecule,
explaining the threshold seen in FIG. 7(b). The threshold force for
restarting the motion among a regular array of harmonic wells, q*E,
is determined by the product of the effective spring constant, k,
and the separation between bases x0: i.e. q*E.about.kx.sub.0. The
trap profile should be invariant for displacing dsDNA by one
basepair, so x.sub.0=0.5*0.34 nm. Thus, the upper bound of the
force required to displace dsDNA between two adjacent traps is 480
pN.
We expect that, in a d=2.0 nm pore, the conformation of the trapped
DNA molecule will resemble that shown in FIG. 8(a) with the
stretching exaggerated by the narrower constriction. Such a trap is
used as part of a sequencing protocol whereby the translocation
kinetics of dsDNA in a pore is stringently controlled and
measurements can then be performed, taking the time necessary to
extract information from the pore current about the identities of
the nucleotides in the constriction. A concern with sequencing
dsDNA in this manner is determining which nucleotide is on which
strand, e.g. distinguishing A-T from T-A. However, our simulations
show that the orientation of the basepair tilt, caused by the
confinement, is maintained during translocation with the
nucleotides of one strand always lagging their partners on the
other. It is, therefore, possible to determine the sequence of
dsDNA without ambiguities between A-T and T-A or G-C and C-G. Two
properties of the trapped dsDNA make sequencing possible. First,
the ions passing through the pore are forced to come in contact
with nucleotides in the constriction. Second, because trapping the
DNA allows current data to be acquired over long time intervals
while the same nucleotides are held near the constriction, the
sequence-dependent current values can be averaged for accuracy. For
sufficiently low biases, the electric potential of the nucleotides
presents an energy barrier to the passage of ions. Because the
passage rate is exponentially related to the height of these
barriers, differences in the heights for different sequences can
have substantial effects on the I-V characteristics.
To demonstrate the feasibility of distinguishing trapped DNA
nucleotides by measuring the ionic current, six systems containing
single T-A, A-T, C-G, G-C, mC-G, and G-mC bps are each simulated
using a combination of MD and Brownian dynamics. Here, X-Y denotes
a system in which a cation passing through the nanopore along the
direction of the electric field encounters nucleotide X first; mC
denotes methylated cytosine. In our setup shown in FIG. 8(b), the
reversing bias is equivalent to replacing an X-Y basepair with Y-X.
FIG. 8(c) shows the absolute values of the current computed from
these simulations. We find that systems containing A and T have
significantly different values of ionic current than those
containing G and C. Moreover, pairs with the same chemical makeup,
but different orientations are distinguishable by the value of the
blocked current (for example, mC-G and G-mC). The accuracy of the
measurement increases as a square root of the number of
translocated ions and, given sufficient averaging time, all
combinations of the nucleotides should be distinguishable. Hence,
our simulations demonstrate the feasibility of distinguishing the
identity of tilted basepairs confined in a nanopore by measuring
the pore current. This is supported by the observation of a signal
attributed to base-pairs slowly translocating through a 2.5 nm pore
shown in FIG. 6(c). The separation in current between A-T and C-G,
which is .DELTA.I=9.4 pA in FIG. 6(b) is smaller than the
prediction (13 pA) for a 2 nm pore, but the trap is also
weaker.
In summary, this example demonstrates sequencing dsDNA in a
nanopore with a diameter smaller than the double helix. The
electric field, in combination with the size of the pore, induce a
stretching transition in the constriction that facilitates control
of both the molecular configuration and the translocation kinetics
at the same time. Thus, high fidelity reads are possible since
control of the molecular configuration (base-pair tilt) determines
how the ions passing through the pore come into contact with the
nucleotides in the constriction, while the translocation kinetics
affect the time interval in which the same nucleotides are held in
the constriction and data is acquired. MD simulations, extrapolated
from our experiments, indicate that to discriminate between A-T and
T-A base-pairs on dsDNA, we demand at minimum that .DELTA.I.about.3
pA for a 2.0 nm pore (in 100 mM KCl). (This signal is not
optimized: e.g. it seems likely that the signal will be larger with
increased electrolyte concentration and a slit geometry).
Therefore, for signal-to-noise >2, we need peak-to-peak noise
<1.5 pA or an rms value of Irms.about.1.5 pA/8=0.2 pA. If
dielectric noise associated with the capacitance predominates, then
where D is the dielectric loss constant (D.about.0.2 for our
membranes). For a bandwidth .DELTA.f.about.1 kHz, we estimate that
DCm.about.1 pF is required to discriminate between base-pairs,
which is a factor 10.times. smaller than the typical membrane
capacitance. On the other hand, a $1000 genome, corresponds to an
estimated throughput of 330,000 bp/sec, which translates to a
capacitance of 10 fF for the same noise current specification. A 10
fF parallel plate capacitance in a 10 nm thick silicon nitride
would have an area of about 1 .mu.m.times.1 .mu.m, which is
accessible with current semiconductor microfabrication technology.
Improvements in the signal may also be achieved using a nano-slit
geometry to force the current closer to the nucleotides by
leveraging recent improvements in TEM technology.
Example 2
Base Discrimination
It is now possible to trap a single molecule of double-stranded DNA
(dsDNA), by stretching it using a nanopore, smaller in diameter
than the double helix, in a solid-state membrane. By applying an
electric force larger than the threshold for stretching, dsDNA can
be impelled through the pore. Once a current blockade associated
with a translocating molecule is detected, the electric field in
the pore is switched in an interval less than the translocation
time to a value below the threshold for stretching. This leaves the
dsDNA stretched in the pore constriction with the base-pairs
tilted, while the B-form canonical structure is preserved outside
the pore. In this configuration, the translocation velocity is
substantially reduced from 1 bp/10 ns to .about.1 bp/2 ms in the
extreme, which facilitates high fidelity reads, allowing us to
discriminate between A-T and C-G base-pairs. Using Molecular
Dynamics simulations to extrapolate to smaller diameters and higher
salt concentration, we find that it is possible to distinguish all
of the trapped dsDNA base-pairs by simply measuring the current.
Further optimizing conditions permits the dsDNA sequence to be
determined without ambiguities between A-T and T-A or G-C and C-G
from the stretched base-pair in the trapped configuration. The
difference between C-G and A-T base-pairs can be resolved in this
case because of the longer (.about.2 ms) time each base-pair spends
in the constriction.
This assertion is corroborated by even longer duration measurements
of blockade currents associated with streptavidin bound, 100 bp
long, C-G and A-T biotinylated duplexes trapped by the electric
field in a pore in a configuration represented schematically in
FIG. 9(a). Streptavidin has an extraordinary affinity for biotin,
which we leverage to measure the blockade current for a trapped
biotin-DNA duplex in a 2.5.times.2.3.+-.0.2 nm pore
(U=0.17.+-.0.04V from .lamda.-DNA blockade frequency) in a
23.1.+-.2.0 nm thick membrane immersed in 1M KCl for transmembrane
biases ranging from 200 mV to 1V. At low voltage, the duration of a
blockade is interminable--the blockade ends only if the voltage is
manually reversed and the dsDNA is impelled out of the pore,
streptavidin and all. However, the dwell time is an exponentially
decreasing function of the applied voltage as illustrated in FIG.
9(b). Nevertheless, the duration of a blockade can still exceed 10
s even at a transmembrane bias of 1V, which is extraordinary (23)
and suggests that the load derived from the electric force on the
trapped DNA may sometimes be shared between the membrane and the
biotin-streptavidin bond. (DNA binding to the membrane could be
attributed to salt-induced absorption on silica in the
nitride(24).) However, neither the dwell time distribution nor the
voltage dependence seems to depend on C-G or A-T variants. The
distribution of dwell times observed at 1V is shown in FIG. 9(c)
for both DNA variants.
In our analysis of the blockades due to the two variants, C-G and
A-T, we focus on dwell times at the peak of the distribution or
longer and on traces with essentially the same open pore current
measured after intervening flushes and cleans. FIGS. 9(d-f) show
typical current blockades for observed at 1V for the peak in the
dwell time distribution, and at 0.8V and 200 mV for dwell times
>10 sec, respectively. Clearly, it is easy to discriminate C-G
base-pairs stretched in the pore from the smaller blockade current
associated with either A-T--the difference is 533.+-.98 pA at a
transmembrane bias of 1V, 789.+-.57 pA at 800 mV and 323.+-.83 pA
at 200 mV. FIG. 9(g) summarizes the differences between the
blockades associated with the two dsDNA variants and open pore
current for the same pore as a function of transmembrane bias. This
summarizes events observed with an open pore current of
I0=2994.+-.51 pA at 200 mV; I0=6134.+-.72 pA at 400 mV; 8174.+-.728
pA at 600 mV; I0=11.909.+-.0.259 nA at 800 mV; and
I0=14.785.+-.0.203 pA at 1V. It seems that C-G can easily be
discriminated from A-T under these conditions over a range of
voltage. Apparently, the increase in molarity (10.times.) and
larger voltage (10.times.) exaggerate the effect of stretching on
the blockade current. This must be the case since larger pore
diameter 3.5.times.2.4.+-.0.2 nm does not reveal a difference
between C-G and A-T beyond for the same salt in the same voltage
range. However, high salt may not be an optimal electrolyte for
sequencing long DNA strands if .lamda.-DNA is absorbed onto the
membrane for 1M KCl and remains trapped in the pore indefinitely
over hrs.
Detailed modeling of DNA translocation: As illustrated in FIG.
4(c), we find that the noise spectrum <100 Hz is sensitive to
the electrolyte concentration while the high frequency (>100 Hz)
noise is not. Hooge suggested that I/f noise occurs in bulk
conductors due to the fluctuating mobility of charge carriers that
produces current fluctuations. In contrast, there are surface
models in which charge traps located on the pore surface have a
fluctuating charge state that affects the ionic current and
likewise exhibits a 1/f characteristic. The two models can be
differentiated by the dependence on the coefficient A on the pore
conductance. FIG. 10 shows the dependence of the coefficient A
measured in a pore with a 2.4.+-.0.2 nm diameter in a 12 nm thick
nitride membrane: the resistance follows a
A.about.R.sup..gamma..sub.p law with .gamma.=1.06.+-.0.15.
Generally, we find that .gamma.=1.03.+-.0.44 over a factor of
10,000.times. in pore resistance, in support of Hooge's
phenomenological picture.
Thus, reducing parasitic capacitances, electrolyte resistance and
amplifier noise are all key elements for improving both the
frequency and noise performance. From the close correspondence
between our models and the measurements of the frequency response,
we assert that this can be accomplished by either: 1. using a
composite membrane consisting of polyimide and silicon nitride
layers to reduce the effect of parasitic elements such as the
depletion layer in the substrate; 2. replacing the silicon handle
wafer altogether with a dielectric substrate; 3. using Ag/AgCl
electrode positioned within microns of the pore; or eliminating the
cable capacitance. For example, FIG. 4(e) explicitly shows the
improvement in the signal to noise ratio that can be gleaned by
using a composite polyimide-nitride membrane. The figure compares a
current blockade associated with a single .lamda.-DNA translocating
through a d=3.0.+-.0.2 nm pore in a 30 nm thick nitride membrane 50
.quadrature.m.times.50 .quadrature.m in area with a pore of the
same diameter in the same type of membrane but with a polyimide
layer 3.6 .quadrature.m thick coating it, reducing the exposed area
to 10 .quadrature.m. We observe a substantial reduction in the
peak-to-peak noise in both the open and blockade pore current (62
pA to 20 pA), which will facilitates signal extraction.
MD confirms our understanding of the mechanism of this process,
indicating: (i) the motion of the dsDNA can be effectively stopped
when the driving voltage is turned off; and (ii) it is feasible to
distinguish trapped DNA base-pairs through simple current
measurements. FIGS. 7 and 11 illustrate two simulated systems,
which include fragments of dsDNA, 100 mM KCl, and pores having 2.6
nm.times.2.1 nm (FIG. 7) and 2.0.times.2.0 nm (FIG. 11)
cross-sections. The molecular conformation within the 2.6
nm.times.2.1 nm pore shown in FIG. 7(a) is stretched by about
8-20%. As illustrated in FIG. 7(b), at 250 and 500 mV, the dsDNA's
motion in the pore is arrested following a small initial
displacement allowed by stretching. Only when a bias of 1V is
applied is dsDNA transport observed. By analyzing dsDNA's
displacements at 0V (shown as inset to FIG. 7(b)), we determine
that the pore acts as a harmonic trap with an effective spring
constant of k=3.0.+-.0.8 nN/nm. A similar analysis for the 2 nm
pore shown in (see FIG. 11) indicates still larger distortions in
the stretched DNA. The base-pair rise increases from 0.34 nm to
0.41 nm at 0V; due to a trap with k=7.2.+-.0.8 nN/nm. (For a d=1.6
nm, the spring constant is even higher: k=19.69.+-.0.06 nN/nm.)
As shown in FIGS. 7(b) and 11(b), MD reveals that a constant
applied voltage of 0.5V is insufficient to restart translocation in
a pore with a diameter smaller than the double helix. However, the
DNA moves in the direction of the applied electric force when 1.0V
is applied. Therefore, a threshold voltage exists between 0.5 and
1.0V. The probability of an escape from a trap depends sharply on
the force applied to the molecule, explaining the thresholds. The
force required to restart the motion is essentially determined by
the product of the spring constant, k, and the separation between
bases x0, which we estimate to be about 0.48 nN, 1.5 nN and 4.1 nN
for the 2.6, 2.0 and 1.6 nm pores respectively.
Based on these observations, we have developed a strategy to
control the translocation and ostensibly to trap bases at fixed
positions within a pore indefinitely. Due to the periodicity of
dsDNA, we find that the translocation through an undersized pore
can proceed in discrete steps in response to a very high-speed
voltage pulse. FIG. 13(e) shows DNA nucleotides near the pore
constriction during and after the application of a half-sine
voltage pulse with a duration of 0.2 ns and amplitudes of 6 and 8V.
For the 6V pulse, the nucleotides are displaced slightly (0-0.2
ns), but return to their original positions after it subsides. On
the other hand, the 8V pulse causes a single base-pair step so that
the final position of the base-pair is near the initial position of
the base-pair directly below it. Thus, we find a tendency for the
portion of the molecule near the constriction to translate in
discrete steps, so that the pore can be used as a turnstile
stepping base-pairs one at a time by applying voltage pulses. These
simulations clearly demonstrate that stepping DNA in a d<2.5 nm
pore is feasible, provided the voltage can be switched fast
enough.
This type of trap process can be used in a sequencing protocol
whereby the translocation kinetics of dsDNA in a pore are
stringently controlled and measurements are performed to extract
the identities of the nucleotides from the pore current. One
difficulty with sequencing this way is determining which nucleotide
is on which strand, e.g. distinguishing A-T from T-A. However, our
in silico experiments show that the base-pair tilt, caused by the
confinement, is maintained during a translocation with the
nucleotides of one strand always lagging their partners on the
other. At low bias, the electric potential of the nucleotides in
the tilted configuration presents a peculiar energy barrier to the
ions with a passage rate that is exponentially related to the
height. The differences in heights for different sequences should
therefore have substantial effects on the current.
To demonstrate the feasibility of distinguishing the nucleotides by
measuring the current, pores containing single T-A, A-T, C-G, and
G-C base-pairs were each simulated as illustrated in FIG. 8. Here,
X-Y denotes a system in which a cation passing through the pore
along the direction of the electric field encounters nucleotide X
first. Due to the symmetry of the setup, a reverse bias is
equivalent to replacing X-Y with Y-X. The table shown in FIG. 8(d)
shows the absolute values of the current computed at salt
concentrations of 100 mM and 1.4M KCl. In the first two rows for
each concentration, we have combined the current values for the
base-pairs containing T and A, and the base-pairs containing C and
G to highlight the fact that these could be clearly discriminated.
We find that the systems containing A and T have significantly
different values of current than those containing G and C.
The simulated current blockades for single, tilted C-G and A-T base
pairs do not match the sequence-dependent blockades measured using
streptavidin-DNA constructs and we don't expect they should. We
attribute this discrepancy to the difference in the conformations
of DNA and to the protein over the pore. In the measurement,
streptavidin anchors a dsDNA molecule in the pore constriction. We
have previously shown the protein in this configuration can
markedly affect the pore current--even causing enhancements above
the open pore value. Moreover, under the influence of the applied
field, dsDNA can stretch in a sequence-dependent way altering the
pore volume available for ion conductance. Sensitive to this
effect, the measurements show a difference between blockade
currents produced by A-T and C-G strands. In contrast, MD shows the
effect of single base-pairs on the ionic current blockade, assuming
that the conformations of the base pairs in the pore are the same.
Through a combination of MD and BD simulations, we expect to
reproduce the sequence-specific ionic blockades measured with
streptavidin.
To obtain a more accurate estimate of the ionic current and allow
for broader exploration of conditions to optimize base-pair
discrimination, we have developed a BD model of the nanopore system
with the interactions derived from all-atom MD simulations. By
modeling water implicitly and using a much larger timestep than can
be used in all-atom MD simulations, the BD simulations show a
10.sup.4-10.sup.5-fold increase in performance. BD simulations of
the pore current have been done before.62-64 However, in contrast
to these studies, advances in computing power allow us to derive
all the parameters of the BD simulations from all-atom MD
simulations. To do this, we make use of umbrella sampling and the
WHAM method to obtain the potential of mean force (PMF) functions
for the interactions. Our BD model takes three types of input: (i)
PMF functions that model the interaction between all pairs of ion
types, (ii) 3D PMF maps that model the environment (including the
solution, the pore, and the DNA) for each type of ion and each DNA
basepair, and (iii) 3D maps of the diffusivity for each type of
ion.
Generating the PMF functions for interactions between the ions via
the WHAM method involved performing many MD simulations with the
distance between the ions restrained to a different value for each
simulation. Likewise, generating the 3D PMF maps required
restraining ions to different positions on a 3D hcp lattice around
the pore and DNA basepair. All these PMF functions implicitly
include the mean effect of water molecules. The diffusivity maps
are computed from these same simulations using the velocity
correlation functions. To validate our BD model, we simulate with
the same parameters used in the all-atom MD simulations above and
compared the ionic currents. For the pore containing no base-pair
at 100 mM KCl we obtain a current of 193.0.+-.0.45 pA for a 96
.mu.s BD simulation, consistent with the MD value. FIG. 12 compares
the pore currents for single base-pairs. For these systems,
including position-dependent diffusivity in the model is crucial to
obtaining quantitative agreement between the BD and MD simulations.
The BD simulations permit us to obtain results in hours that we
could not obtain in weeks of all-atom MD simulations. As shown in
FIG. 12, we have demonstrated the feasibility of distinguishing
base-pairs of the same chemical structure but different
orientations, T-A can be easily distinguished from A-T and C-G can
be readily distinguished from G-C.
Finally, FIG. 13 illustrates the coincidence between this naive
model of the pore current derived from the MD results of FIG. 12
for a single base-pair trapped in the constriction and the
measurements obtained near 58.8 sec, just before the molecule exits
the pore, assuming a uniform translocation velocity of 1 bp/2 ms.
The current trace is extracted at the end of the blockade, which
should amount to approximately the last 50 bps, and then compared
with the corresponding sequence (from 48453-48502 bp) associated
with the 3' end of .lamda.-DNA. There is an offset in the current
between the two traces of 117 pA, since neither the voltage nor the
pore diameter used to extract the values used in the model coincide
with the measurement. The correlation between the measurement and
simulation is 0.31, which potentially indicates a lack of
signal-to-noise required for sequencing with single base resolution
or a non-uniform translocation velocity. However, this
correspondence indicates the prospects for sequencing with single
base resolution and provides a target to exceed through
optimization of the pore geometry and electrolyte.
Optimization of the Pore and Electrolyte for Sequencing.
While both simulation and measurements indicate high electrolyte
concentration for improved signal fidelity, high salt may not be an
optimal for sequencing long DNA strands. We observed that at high
salt concentration (1M KCl) .lamda.-DNA adheres to the pore even at
high transmembrane bias; apparently it is absorbed onto the
membrane and it remains there for hours. A typical example is
illustrated in FIG. 13(a) with a constant 1V transmembrane bias
applied. While short duration (with a typical duration of
.about.6.8 ms) current blockades with .DELTA.I/I0.about.0.36 can be
observed in the interval from 0-0.5 s, we also find an extended
blockade event with .DELTA.I/I0>0.62, starting near 0.6 s, that
persists interminably. We attribute the blockade to one or more
dsDNA bound to the membrane in the pore. To alleviate this sticking
we can use shorter strands or lower molarity electrolyte, higher or
lower pH, and thinner membranes. On the other hand, the sticking
frequency seems to increase with the DNA concentration, the salt
concentration and if a divalent electrolyte such as CaCl2 is used
in place of KCl.
Salt-induced absorption on silica is one of the most common methods
for extracting DNA from cell homogenates and it is consistent with
the observations we have made about the sticking conditions in a
pore in a nitride membrane: i.e. DNA sticks at high molarity, but
not at dilute concentrations. Although the mechanism is not
understood, it is supposed that a high electrolyte concentration
disrupts the shell of hydration around the DNA allowing a
positively charged ion to form a salt bridge between the negatively
charged silica and the negatively charged DNA backbone. We
attribute the observation of the interminably trapped dsDNA to
binding to oxide in a pore that is likely formed after sputtering
exposure to air and/or electrolyte or surface treatments such as a
O2 plasma or piranha clean. After plasma cleaning, CVD silicon
nitride films seem to show high oxygen to silicon ratio (0.55),
indicating severe oxidation, while the carbon content decreases and
the silicon to nitrogen ratio increases (from 0.73 to 1.23.)
according to XPS.
FIGS. 13(b,c) show a test of single molecule fluorescence obtained
from DNA duplexes intercalated with YOYO-1 (a dimeric cyanine
fluorescent dye) fluorochromes adsorbed onto a silanized silica
coverslip and silicon nitride membrane in 1M KCl. The DNA molecules
are first stained with YOYO 1 at a ratio of DNA base pairs to
YOYO-1 molecules of 10:1 and diluted to 1.67 nM (in 1M KCl).
Subsequently, the coverslips were cleaned using an O2 plasma (we
also tested ethanol and H2SO4:H2O2 treatments and saw no
difference) and then bonded them with a microfluidic, and
subsequently immersed it a 1M KCl electrolyte containing the DNA.
YOYO emits at 510 nm when excited with 457 nm wavelength. The
fluorescence was observed using a Zeiss Axio-Observer inverted
microscope with an Achroplan 100.times. oil immersion objective
(1.3NA) equipped with a 1024.times.10.sup.24 EMCCD camera (Andor,
DU-888) operated at -70.degree. C. The fluorescence channel is
defined by excitation filter 438/24 nm center wavelength/bandwidth,
dichroic mirror with 458 nm edge wavelength, emission filter at
emission 483/32 nm. The excitation is provided by EXFO X-Cite 120
metal halide lamp operated at 12% iris. We clearly observe DNA
sticking to the silica in FIG. 13(b), while we find a dearth of DNA
sticking to the nitride membrane (the fluorescent DNA can be seen
hovering near positions indicated by the arrows) treated with the
same conditions as shown in FIG. 13(c). These data
counter-indicates the use of high salt and silicon oxide surfaces
for sequencing long DNA strands, and while nitride membrane seems
more optimal than oxide, the concentration of dsDNA and electrolyte
have to be optimized to avoid absorption in the pore where the
surface to volume ratio is very high while still providing adequate
signal.
STATEMENTS REGARDING INCORPORATION BY REFERENCE AND VARIATIONS
All references throughout this application, for example patent
documents including issued or granted patents or equivalents;
Patent application publications; and non-patent literature
documents or other source material; are hereby incorporated by
reference herein in their entireties, as though individually
incorporated by reference, to the extent each reference is at least
partially not inconsistent with the disclosure in this application
(for example, a reference that is partially inconsistent is
incorporated by reference except for the partially inconsistent
portion of the reference). This application is related to U.S.
Provisional Patent Application No. 61/139,056 filed Dec. 19, 2008
titled "Detecting and Sorting Methylated DNA using a Synthetic
Nanopore", from which PCT Pub. No. WO 2010/080617 (GS 168-08)
claims benefit, each of which is hereby incorporated by reference
to the extent not inconsistent herewith.
The terms and expressions which have been employed herein are used
as terms of description and not of limitation, and there is no
intention in the use of such terms and expressions of excluding any
equivalents of the features shown and described or portions
thereof, but it is recognized that various modifications are
possible within the scope of the invention claimed. Thus, it should
be understood that although the present invention has been
specifically disclosed by preferred embodiments, exemplary
embodiments and optional features, modification and variation of
the concepts herein disclosed may be resorted to by those skilled
in the art, and that such modifications and variations are
considered to be within the scope of this invention as defined by
the appended claims. The specific embodiments provided herein are
examples of useful embodiments of the present invention and it will
be apparent to one skilled in the art that the present invention
may be carried out using a large number of variations of the
devices, device components, methods steps set forth in the present
description. As will be obvious to one of skill in the art, methods
and devices useful for the present methods can include a large
number of optional composition and processing elements and
steps.
When a group of substituents is disclosed herein, it is understood
that all individual members of that group and all subgroups, are
disclosed separately. When a Markush group or other grouping is
used herein, all individual members of the group and all
combinations and subcombinations possible of the group are intended
to be individually included in the disclosure.
Every formulation or combination of components described or
exemplified herein can be used to practice the invention, unless
otherwise stated.
Whenever a range is given in the specification, for example, a
temperature range, a size or distance range, a time range, or a
composition or concentration range, all intermediate ranges and
subranges, as well as all individual values included in the ranges
given are intended to be included in the disclosure. It will be
understood that any subranges or individual values in a range or
subrange that are included in the description herein can be
excluded from the claims herein.
All patents and publications mentioned in the specification are
indicative of the levels of skill of those skilled in the art to
which the invention pertains. References cited herein are
incorporated by reference herein in their entirety to indicate the
state of the art as of their publication or filing date and it is
intended that this information can be employed herein, if needed,
to exclude specific embodiments that are in the prior art. For
example, when composition of matter are claimed, it should be
understood that compounds known and available in the art prior to
Applicant's invention, including compounds for which an enabling
disclosure is provided in the references cited herein, are not
intended to be included in the composition of matter claims
herein.
As used herein, "comprising" is synonymous with "including,"
"containing," or "characterized by," and is inclusive or open-ended
and does not exclude additional, unrecited elements or method
steps. As used herein, "consisting of" excludes any element, step,
or ingredient not specified in the claim element. As used herein,
"consisting essentially of" does not exclude materials or steps
that do not materially affect the basic and novel characteristics
of the claim. In each instance herein any of the terms
"comprising", "consisting essentially of" and "consisting of" may
be replaced with either of the other two terms. The invention
illustratively described herein suitably may be practiced in the
absence of any element or elements, limitation or limitations which
is not specifically disclosed herein.
All art-known functional equivalents, of any such materials and
methods are intended to be included in this invention. The terms
and expressions which have been employed are used as terms of
description and not of limitation, and there is no intention that
in the use of such terms and expressions of excluding any
equivalents of the features shown and described or portions
thereof, but it is recognized that various modifications are
possible within the scope of the invention claimed. Thus, it should
be understood that although the present invention has been
specifically disclosed by preferred embodiments and optional
features, modification and variation of the concepts herein
disclosed may be resorted to by those skilled in the art, and that
such modifications and variations are considered to be within the
scope of this invention as defined by the appended claims.
TABLE-US-00001 TABLE 1 Selected parameters for the model shown in
FIG. 3(h) used to fit the measurements of three membranes shown in
FIG. 3(g). Imide/ 12 nm nitride 30 nm nitride 200 nm nitride Model
parameters 50 .mu.m .times. 50 .mu.m 10 .mu.m .times. 10 .mu.m 500
.mu.m .times. 500 .mu.m Rel Electrolyte Res. (k.OMEGA.) 0.85 0.61
0.9 Fdlt1 Top double layer Faradaic 2.80 .times. 10.sup.7 7.30
.times. 10.sup.6 4.00 .times. 10.sup.6 Coeff.(k.OMEGA.) Cdlt1 Top
double layer Cap. (pF) 303000 303000 184000 Rim1 Polyimide Res.
(G.OMEGA.) n/a 122 n/a Cim1 Polyimide Cap. (pF) n/a 18.3 n/a Rmem1
Si3N4 Res. (G.OMEGA.) 795 1600 3890 Cmem1 Si3N4 Cap. (pF) 1520 727
150 Rdt Top depletion region Res. (k.OMEGA.) 22.7 22.7 91.2 Cdt Top
depletion region Cap. (pF) 308 308 46.7 Rsi Si Res. (.OMEGA.) 25 25
21.8 Rdb Bottom depletion region Res. (k.OMEGA.) 706 706 800 Cdb
Bottom depletion region Cap. (pF) 895 399 300 Rim2 Polyimide Res.
(G.OMEGA.) n/a 1410 n/a Cim2 Polyimide Cap. (pF) n/a 0.007 n/a
Rmem2 Si3N4 Res. (G.OMEGA.) n/a 15000 n/a Cmem2 Si3N4 Cap. (pF) n/a
3.7 n/a Cmem3 Si3N4 Cap. (pF) 8.1 0.08 169
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